Textbook of Reumathology-Kelley's PDF [PDF]

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VOLUME



I



KELLEY’S



Textbook of Rheumatology EIGHTH EDITION



Gary S. Firestein, MD



Ralph C. Budd, MD



Edward D. Harris, Jr., MD



Iain B. McInnes, PhD, FRCP



Shaun Ruddy, MD



John S. Sergent, MD



Professor of Medicine Chief, Division of Rheumatology, Allergy, and Immunology Dean, Translational Medicine University of California, San Diego, School of Medicine La Jolla, California Professor of Medicine Director, Immunobiology Program University of Vermont College of Medicine Burlington, Vermont George DeForest Barnett Professor of Medicine, Emeritus Stanford University School of Medicine Academic Secretary to Stanford University, Emeritus Stanford University Stanford, California Professor of Experimental Medicine Honorary Consultant Rheumatologist Centre for Rheumatic Diseases, Faculty of Medicine University of Glasgow Glasgow, United Kingdom Professor Emeritus, Department of Internal Medicine, Division of Rheumatology, Allergy, and Immunology Virginia Commonwealth University School of Medicine at the Medical College of Virginia Richmond, Virginia Professor of Medicine Vice Chair for Education and Residency Program Director Vanderbilt University School of Medicine Nashville, Tennessee



1600 John F. Kennedy Blvd. Ste 1800 Philadelphia, PA 19103-2899



ISBN: 978-1-4160-3285-4 (Expert Consult) KELLEY’S TEXTBOOK OF RHEUMATOLOGY 978-1-4160-4842-8 (Expert Consult Premium Ed.) Copyright © 2009, 2005, 2001, 1997, 1993, 1989, 1985, 1981 by Saunders, an imprint of Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Permissions may be sought directly from Elsevier’s Rights Department: phone: (+1) 215 239 3804 (US) or (+44) 1865 843830 (UK); fax: (+44) 1865 853333; e-mail: [email protected]. You may also complete your request on-line via the Elsevier website at http://www.elsevier.com/permissions.



Notice Knowledge and best practice in this field are constantly changing. As new research and experience broaden our knowledge, changes in practice, treatment, and drug therapy may become necessary or appropriate. Readers are advised to check the most current information provided (i) on procedures featured or (ii) by the manufacturer of each product to be administered, to verify the recommended dose or formula, the method and duration of administration, and contraindications. It is the responsibility of the practitioner, relying on experience and knowledge of the patient, to make diagnoses, to determine dosages and the best treatment for each individual patient, and to take all appropriate safety precautions. To the fullest extent of the law, neither the Publisher nor the Editors assume any liability for any injury and/or damage to persons or property arising out of or related to any use of the material contained in this book. The Publisher



Library of Congress Cataloging-in-Publication Data Kelley’s textbook of rheumatology / [edited by] Gary S. Firestein ... [et al.]. -- 8th ed. p. ; cm. Includes bibliographical references and index. ISBN 978-1-4160-3285-4 1. Rheumatology. 2. Rheumatism. 3. Arthritis. I. Firestein, Gary S. II. Kelley, William N., 1939III. Title: Textbook of rheumatology. [DNLM: 1. Rheumatic Diseases. 2. Arthritis. WE 544 K29 2009] RC927.T49 2009 616.7'23--dc22 2007048387



Acquisitions Editor: Kimberly Murphy Developmental Manager: Cathy Carroll Developmental Editor: Angela Norton Publishing Services Manager: Linda Van Pelt Project Manager: Francisco Morales Design Direction: Ellen Zanolle Cover Design: Ellen Zanolle



Printed in Canada Last digit is the print number: 9



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Sincerest thanks to my wonderful wife, Linda, and our children, David and Cathy, for their patience and support. Also, the editorial help of our two Cavalier King Charles puppies, Winston and Humphrey, was invaluable. Gary S. Firestein



Sincere thanks for the kind mentoring from Edward D. Harris Jr., H. Robson MacDonald, and C. Garrison Fathman, as well as for the support of my wife, Lenore, and my children, Graham and Laura. Ralph C. Budd



Many thanks to my mentor, Steve Krane, and for the support of the Harris boys, Ned, Tom, and Chandler, and Eileen . . . and for the happy smiles of the grandkids— Andrew, Eliza, Maeve, and Liam. Ted Harris



To my wife, Karin, for her patience, understanding, and love and to our wonderful girls, Megan and Rebecca, who continue to enlighten me. Iain B. McInnes



To my wife, Millie; our children, Christi and Candace; and our grandchildren, Kevin, Matthew, and Katharine. Shaun Ruddy



To Carole and our children, Ellen and Katie, and to our grandchildren, Kathryn, Henry, Emmaline, and Romy. John S. Sergent



CONTRIBUTORS



Steven B. Abramson, MD Professor of Medicine and Pathology New York University School of Medicine New York, New York   Neutrophils and Eosinophils; Pathogenesis of Osteoarthritis Leyla Alparslan, MD Instructor in Orthopaedic Radiology Uppsala University Faculty of Medicine Staff Radiologist, Akademiska Hospital Uppsala, Sweden   Imaging Modalities in Rheumatic Disease Thomas P. Andriacchi, PhD Professor Department of Mechanical Engineering Stanford University School of Engineering Department of Orthopaedics Stanford University School of Medicine Stanford VA Palo Alto Research & Development, Bone and Joint Research Center Palo Alto, California  Joint Biomechanics: The Role of Mechanics in Joint Pathology John P. Atkinson, MD Samuel B. Grant Professor of Medicine and Professor of Molecular Microbiology Washington University in St. Louis School of Medicine Physician, Barnes-Jewish Hospital St. Louis, Missouri   Complement System Stefan Bachmann, MD FMH Specialist in Internal Medicine and Rheumatology FMH Specialist in Physical Medicine and Rehabilitation Leitender Artz/Chefarzt-Stellvertreter Klinik für Rheumatologie und Rehabilitation des Bewegungsapparates Valens, Switzerland   Introduction to Physical Medicine and Rehabilitation Leslie R. Ballou, PhD Professor of Medicine and Molecular Sciences, Department of Rheumatology University of Tennessee College of Medicine Research Chemist, VA Medical Center and UT Health Science Center Memphis, Tennessee   Nonsteroidal Anti-inflammatory Drugs



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Stanley P. Ballou, MD Associate Professor of Medicine Case Western Reserve University School of Medicine Director of Rheumatology MetroHealth Medical Center Cleveland, Ohio   Acute-Phase Reactants and the Concept of Inflammation Walter G. Barr, MD Professor of Medicine Northwestern University Feinberg School of Medicine Chicago, Illinois   Mycobacterial Infections of Bones and Joints; Fungal Infections of the Bones and Joints Dorcas Eleanor Beaton, BScOT, MSc, PhD Assistant Professor, Department of Health Policy, Management and Evaluation University of Toronto Faculty of Medicine Scientist and Director, Mobility Program Clinical Research Unit St. Michael’s Hospital Toronto, Ontario, Canada   Assessment of Health Outcomes Robert M. Bennett, MD, FRCP, MACR Professor of Medicine and Nursing Research Oregon Health & Science University School of Medicine and School of Nursing Portland, Oregon   Overlap Syndromes Francis Berenbaum, MD, PhD Professor of Rheumatology Pierre and Marie Curie University (UPMC—Paris Universitas) Faculty of Medicine Hospital Saint-Antoine Paris, France   Clinical Features of Osteoarthritis Johannes W.J. Bijlsma, MD, PhD Professor and Chair, Department of Rheumatology   & Clinical Immunology University Medical Center Utrecht Utrecht, The Netherlands   Glucocorticoid Therapy Linda K. Bockenstedt, MD Harold W. Jockers Professor of Medicine Department of Internal Medicine, Section of Rheumatology Yale University School of Medicine New Haven, Connecticut   Lyme Disease



CONTRIBUTORS



Maarten Boers, MSc, MD, PhD Professor of Clinical Epidemiology Department of Clinical Epidemiology and Biostatistics VU University Amsterdam Faculty of Medicine Amsterdam, The Netherlands   Assessment of Health Outcomes Robert Alan Bonakdar, MD Assistant Clinical Professor, Department of Family and Preventive Medicine UC San Diego School of Medicine Director of Pain Management Scripps Center for Integrative Medicine La Jolla, California   Integrative Medicine in Rheumatology:   An Evidence-Based Approach Dimitrios T. Boumpas, MD, FACP Professor and Chairman, Department of Internal Medicine, Division of Rheumatology, Clinical Immunology, and Allergy University of Crete Medical School Chief of Medicine Heraklion University General Hospital Crete, Greece   Clinical Features and Treatment of Systemic Lupus Erythematosus Barry Bresnihan, MD Professor of Rheumatology University College Dublin School of Medicine   and Medical Science National University of Ireland Consultant Rheumatologist St. Vincent’s University Hospital Prinicipal Investigator Conway Institute of Biomedical Research Dublin, Ireland   Synovium Doreen B. Brettler, MD Professor of Medicine University of Massachusetts Medical School Director, New England Hemophilia Center University of Massachusetts Memorial Healthcare Worcester, Massachusetts   Hemophilic Arthropathy Paul L. Briant, PhD, MS VA Palo Alto Research & Development Bone and Joint Research Center Palo Alto, CA   Joint Biomechanics: The Role of Mechanics in Joint Pathology Ralph C. Budd, MD Professor of Medicine Director, Immunobiology Program University of Vermont College of Medicine Burlington, Vermont   T Lymphocytes



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Leonard H. Calabrese, DO Professor of Medicine and R.J. Fasenmyer Chair of Clinical Immunology Cleveland Clinic Lerner College of Medicine Vice Chairman, Department of Rheumatic and Immunologic Diseases Cleveland Clinic Foundation Cleveland, Ohio   Antineutrophil Cytoplasmic Antibody–Associated Vasculitis Amy C. Cannella, MD Assistant Professor, Department of Medicine, Section of Rheumatology and Immunology University of Nebraska College of Medicine Omaha, Nebraska   Methotrexate, Leflunomide, Sulfasalazine, Hydroxychloroquine, and Combination Therapies Eugene J. Carragee, MD Professor of Orthopaedic Surgery Stanford University School of Medicine Director, Spine Surgery Section Stanford University Hospital and Clinics Stanford, California   Low Back Pain Steven Carsons, MD Professor of Medicine State University of New York at Stony Brook School   of Medicine Stony Brook Chief, Division of Rheumatology, Allergy   and Immunology Winthrop University Hospital Mineola, New York   Sjögren’s Syndrome James T. Cassidy, MD Professor, Department of Child Health University of Missouri–Columbia School of Medicine Chief of Pediatric Rheumatology University of Missouri Health Sciences Center Columbia, Missouri   Systemic Lupus Erythematosus, Juvenile Dermatomyositis, Scleroderma, and Vasculitis Eliza F. Chakravarty, MD, MS Assistant Professor, Department of Medicine, Division   of Immunology and Rheumatology Stanford University School of Medicine Stanford, California   Musculoskeletal Syndromes in Malignancy Christopher Chang, MD, PhD Associate Clinical Professor, Department of Internal Medicine, Division of Rheumatology/Allergy/Clinical Immunology UC Davis School of Medicine Sacramento Staff, UC Davis Genome and Biomedical Services   Facility Davis, California   Osteonecrosis



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CONTRIBUTORS



Joseph S. Cheng, MD, MS Assistant Professor of Neurological Surgery Vanderbilt University School of Medicine Director, Neurosurgery Spine Program Vanderbilt University Medical Center Nashville, Tennessee   Neck Pain



Jeroen DeGroot, MD Operations Manager, Inflammatory and Degenerative Diseases BioSciences Division TNO Quality of Life Leiden, The Netherlands   Biologic Markers



Christopher P. Chiodo, MD Instructor in Orthopaedic Surgery, Department   of Orthopedic Surgery Harvard Medical School Chief, Foot and Ankle Division Brigham and Women’s Hospital Boston, Massachusetts   Foot and Ankle Pain



Christopher P. Denton, PhD, FRCP Professor of Experimental Rheumatology Royal Free and University College Medical School Honorary Consultant Rheumatologist, Centre for Rheumatology London, United Kingdom   Systemic Sclerosis and the Scleroderma-Spectrum Disorders



Paul P. Cook, MD Associate Professor of Medicine Division of Infectious Diseases Department of Infectious Diseases Department of Internal Medicine Brody School of Medicine at East Carolina University Greenville, North Carolina   Bacterial Arthritis



Clinton Devin, MD Orthopedic Surgeon, Department of Orthopaedics   and Rehabilitation Vanderbilt Sports Medicine Center Nashville, Tennessee   Neck Pain



Joseph E. Craft, MD Professor of Medicine and Immunobiology Chief, Section of Rheumatology, and Director, Investigative Medicine Yale University School of Medicine Chief of Rheumatology and Attending Physician Yale–New Haven Hospital New Haven, Connecticut   Antinuclear Antibodies Gaye Cunnane, MD, MB, PhD, FRCPI Senior Lecturer in Medicine Trinity College Dublin Faculty of Health Sciences School of Medicine Consultant in Rheumatology and Internal Medicine St. James’ Hospital Dublin, Ireland   Hemochromatosis Jody A. Dantzig, BS, PhD Medical Student University of Pennsylvania School of Medicine Philadelphia, Pennsylvania   Muscle: Anatomy, Physiology, and Biochemistry John M. Davis III, MD Assistant Professor of Medicine Division of Rheumatology Mayo Clinic Rochester, Minnesota   History and Physical Examination of the Musculoskeletal System



Betty Diamond, MD Professor, Department of Microbiology & Immunology   and Department of Medicine (Rheumatology) Albert Einstein College of Medicine Bronx Head and Investigator, Center for Autoimmune   and Musculoskeletal Diseases The Feinstein Institute for Medical Research Manhasset, New York   B Cells Federico Díaz-González, MD Associate Professor of Rheumatology Universidad de La Laguna Faculty of Medicine Staff Rheumatologist Hospital Universitario de Canarias La Laguna, Spain   Platelets and Rheumatic Diseases Paul E. Di Cesare, MD, FACS Professor and Michael W. Chapman Chair, Department   of Orthopaedic Surgery UC Davis School of Medicine Sacramento, California   Pathogenesis of Osteoarthritis Joost P.H. Drenth, MD, PhD Professor of Molecular Gastroenterology and Hepatology Department of Gastroenterology and Hepatology Radboud University Nijmegen Medical Centre Faculty of Medical Sciences Nijmegen, The Netherlands   Familial Auto-inflammatory Syndromes George F. Duna, MD, FACP Associate Professor of Medicine Baylor College of Medicine Houston, Texas   Antineutrophil Cytoplasmic Antibody–Associated Vasculitis



CONTRIBUTORS



Michael L. Dustin, PhD Irene Diamond Professor of Immunology and Associate Professor of Pathology Department of Molecular Pathogenesis The Helen L. and Martin S. Kimmel Center for Biology and Medicine, Skirball Institute of Biomolecular Medicine New York University School of Medicine New York, New York   Adaptive Immunity Including Organization of Lymphoid Tissues Hani S. El-Gabalawy, MD, FRCPC Professor of Medicine and Immunology and Head, Division of Rheumatology University of Manitoba Faculty of Medicine Rheumatologist Winnipeg Health Sciences Centre Winnipeg, Manitoba, Canada   Synovial Fluid Analysis, Synovial Biopsy, and Synovial Pathology Keith B. Elkon, MD Professor of Medicine and Immunology and Head, Division of Rheumatology Department of Medicine University of Washington School of Medicine Seattle, Washington   Cell Survival and Death in Rheumatic Diseases Doruk Erkan, MD Assistant Professor of Medicine Weill Medical College of Cornell University Associate Physician-Scientist and Assistant Attending Physician Barbara Volcker Center for Women and Rheumatic Diseases Hospital for Special Surgery New York, New York   Antiphospholipid Syndrome Gary S. Firestein, MD Professor of Medicine Chief, Division of Rheumatology, Allergy,   and Immunology Dean, Translational Medicine University of California, San Diego, School of Medicine La Jolla, California   Etiology and Pathogenesis of Rheumatoid Arthritis; Clinical Features of Rheumatoid Arthritis Oliver FitzGerald, MD, FRCPI, FRCP(UK) Newman Clinical Research Professor University College Dublin School of Medicine   and Medical Science National University of Ireland Consultant Rheumatologist St. Vincent’s University Hospital Dublin, Ireland   Psoriatic Arthritis



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John P. Flaherty, MD Professor of Medicine Associate Chief and Director of Clinical Services, Division of Infectious Diseases Northwestern University Feinberg School of Medicine Chicago, Illinois   Mycobacterial Infections of Bones and Joints; Fungal Infections of the Bones and Joints Adrienne M. Flanagan, MD, PhD Professor Institute of Orthopaedics and Musculoskeletal Science University College London London Royal National Orthopaedic Hospital Stanmore Department of Histopathology, University College Hospital London, United Kingdom   Synovium Karen A. Fortner, PhD Research Assistant Professor Immunobiology Program Department of Medicine University of Vermont College of Medicine Burlington, Vermont   T Lymphocytes Howard A. Fuchs, MD Associate Professor of Medicine Division of Rheumatology Vanderbilt University Tennessee Valley Healthcare System Department of Veterans Affairs Medical Center Nashville, TN   Polyarticular Arthritis Steffen Gay, MD Professor Department of Rheumatology University Hospital Zurich, Switzerland   Fibroblasts and Fibroblast-like Synoviocytes Mark C. Genovese, MD Professor of Medicine Stanford University School of Medicine Co-Chief, Division of Immunology and Rheumatology Stanford University Medical Center Stanford, California   Treatment of Rheumatoid Arthritis M. Eric Gershwin, MD Distinguished Professor of Medicine UC Davis School of Medicine Sacramento, California   Osteonecrosis



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CONTRIBUTORS



Allan Gibofsky, MD, JD, FACP, FCLM Professor of Medicine and Public Health Weill Medical College of Cornell University Adjunct Professor of Law Fordham University School of Law Attending Rheumatologist Hospital for Special Surgery New York, New York   Poststreptoccocal Arthritis and Rheumatic Fever Mark H. Ginsberg, MD Professor, Department of Medicine, Rheumatology Section UC San Diego School of Medicine La Jolla, California   Platelets and Rheumatic Diseases Joseph Golbus, MD Associate Professor of Medicine Northwestern University Feinberg School of Medicine Senior Attending Physician, Division of Rheumatology President Evanston Northwestern Healthcare Medical Group Chicago, Illinois   Monarticular Arthritis Yale E. Goldman, MD, PhD Professor, Department of Physiology University of Pennyslvania School of Medicine Director, Pennsylvania Muscle Institute Philadelphia, Pennsylvania   Muscle: Anatomy, Physiology, and Biochemistry Mary B. Goldring, PhD Weill Medical College of Cornell University Senior Scientist Hospital for Special Surgery New York, New York   Biology of the Normal Joint; Cartilage and Chondrocytes Steven R. Goldring, MD Weill Medical College of Cornell University Chief Scientific Officer Hospital for Special Surgery New York, New York   Biology of the Normal Joint Stuart B. Goodman, MD, PhD, FRCSC, FACS, FBSE Robert L. and Mary Ellenburg Professor of Surgery, Department of Orthopaedic Surgery Stanford University School of Medicine Attending Orthopaedic Surgeon, Stanford University Medical Center Consultant Orthopaedic Surgeon, Lucile Salter Packard Children’s Hospital at Stanford Stanford Consultant Orthopaedic Surgeon, Palo Alto Veterans Administration Hospital Palo Alto, California   Hip and Knee Pain



Carl S. Goodyear, PhD Lecturer and Arthritis Research Campaign University of Glasgow Faculty of Medicine NCCD Fellow, Division of Clinical Neurosciences Glasgow Biomedical Research Centre Glasgow, United Kingdom   Rheumatoid Factors and Other Autoantibodies in Rheumatoid Arthritis Siamon Gordon, MBChB, PhD, FRS, FMedSci Professor Emeritus Sir William Dunn School of Pathology University of Oxford Oxford, United Kingdom   Mononuclear Phagocytes in Rheumatic Diseases Adam Greenspan, MD, FACR Professor Emeritus of Radiology Department of Radiology, Section of Musculoskeletal Imaging UC Davis School of Medicine Sacramento, California Osteonecrosis Peter K. Gregersen, MD Professor of Medicine and Pathology New York University School of Medicine New York, New York   Genetics of Rheumatic Diseases Christine Grimaldi, PhD Assistant Professor, Department of Microbiology & Immunology Albert Einstein College of Medicine Bronx Assistant Investigator, Center for Autoimmune and Musculoskeletal Disease The Feinstein Institute for Medical Research Manhasset, New York   B Cells Bevra Hannahs Hahn, MD, FACR, MACR Professor and Vice Chair, Department of Medicine David Geffen School of Medicine at UCLA Chief, Rheumatology and Arthritis UCLA Medical Center Los Angeles, California   Pathogenesis of Systemic Lupus Erythematosus J. Timothy Harrington, MD Associate Professor, Department of Medicine University of Wisconsin School of Medicine and Public Health Madison, Wisconsin   Mycobacterial Infections of Bones and Joints; Fungal Infections of the Bones and Joints Edward D. Harris, Jr., MD, MACR George DeForest Barnett Professor of Medicine, Emeritus Stanford University School of Medicine Academic Secretary to Stanford University, Emeritus Stanford University Stanford, California   Clinical Features of Rheumatoid Arthritis



CONTRIBUTORS



David B. Hellmann, MD Aliki Perroti Professor of Medicine Johns Hopkins University School of Medicine Vice Dean and Chairman, Department of Medicine Johns Hopkins Bayview Medical Center Baltimore, Maryland   Giant Cell Arteritis, Polymyalgia Rheumatica,   and Takayasu’s Arteritis George Ho, Jr., MD Professor of Medicine Brody School of Medicine at East Carolina University Greenville, North Carolina   Bacterial Arthritis James I. Huddleston, MD Assistant Professor, Department of Orthopaedic Surgery Stanford University School of Medicine Stanford, California   Hip and Knee Pain Gene G. Hunder, MD Professor Emeritus Mayo Clinic College of Medical Sciences Emeritus Member Department of Internal Medicine, Division of Rheumatology Mayo Clinic Rochester, Minnesota   History and Physical Examination of the Musculoskeletal System



Arthur Kavanaugh, MD Professor of Medicine Center for Innovative Therapy, Division of Rheumatology, Allergy and Immunology UC San Diego School of Medicine La Jolla, California   Anticytokine Therapies Alisa E. Koch, MD Frederick G.L. Huetwell and William D. Robinson, MD Professor of Rheumatology University of Michigan Medical School Ann Arbor, Michigan   Cell Recruitment and Angiogenesis Deborah Krakow, MD Associate Professor of Obstetrics and Gynecology David Geffen School of Medicine at UCLA Attending Physician, Department of Obstetrics   and Gynecology Cedars-Sinai Medical Center Los Angeles, California   Heritable Diseases of Connective Tissue Joel M. Kremer, MD Pfaff Family Professor of Medicine Albany Medical College Director of Research The Center for Rheumatology Albany, New York   Nutrition and Rheumatic Diseases



Johannes W.G. Jacobs, MD, PhD Associate Professor, Department of Rheumatology   and Clinical Immunology Rheumatologist and Senior Researcher University Medical Center Utrecht Utrecht, The Netherlands   Glucocorticoid Therapy



Hollis E. Krug, BS, MD Associate Clinical Professor of Medicine University of Minnesota Medical School Staff Rheumatologist VA Medical Center Minneapolis, Minnesota   Management of Chronic Pain



Joanne M. Jordan, MD, MPH Associate Professor of Medicine and Orthopaedics Chief, Division of Rheumatology, Allergy, and Immunology University of North Carolina at Chapel Hill School   of Medicine Director, Thurston Arthritis Research Center Chapel Hill, North Carolina   Principles of Epidemiology in Rheumatic Disease



Irving Kushner, MD Professor of Medicine Case Western Reserve University School of Medicine Staff Rheumatologist MetroHealth Medical Center Cleveland, Ohio   Acute-Phase Reactants and the Concept of Inflammation



Joseph L. Jorizzo, MD Professor and Former (Founding) Chair, Department   of Dermatology Wake Forest University School of Medicine Winston-Salem, North Carolina   Behçet’s Disease Kenneth C. Kalunian, MD Professor of Medicine and Director of Rheumatology, Allergy and Immunology UC San Diego School of Medicine La Jolla, California   Rheumatic Manifestations of Hemoglobinopathies



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Robert B.M. Landewé, MD Professor of Rheumatology, Department of Internal Medicine, Division of Rheumatology Maastricht University Faculty of Medicine Staff Rheumatologist Atrium Medical Center Heerlen Maastricht, The Netherlands   Clinical Trial Design and Analysis Nancy E. Lane, MD Professor of Medicine and Rheumatology UC Davis School of Medicine Director, Center for Healthy Aging Sacramento, California   Metabolic Bone Disease



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CONTRIBUTORS



Daniel J. Laskin, DDS, MS Professor and Chairman Emeritus, Department of Oral   and Maxillofacial Surgery School of Dentistry and School of Medicine Virginia Commonwealth University Richmond, Virginia   Temporomandibular Joint Pain David M. Lee, MD Assistant Professor of Medicine Harvard Medical School Associate Physician Brigham and Women’s Hospital Boston, Massachusetts   Mast Cells Lela A. Lee, MD Professor of Dermatology and Medicine University of Colorado School of   Medicine Director of Dermatology Denver Health Medical Center Denver, Colorado   The Skin and Rheumatic Diseases Marjatta Leirisalo-Repo, MD, PhD Professor of Rheumatology, Department of Medicine, Division of Rheumatology Helsinki University Faculty of Medicine Staff Rheumatologist, Helsinki University Central Hospital Helsinki, Finland   Undifferentiated Spondyloarthritis and Reactive Arthritis David C. Leopold, MD, DABFM Faculty Physician and Director, Integrative Medical Education Scripps Center for Integrative Medicine La Jolla, California   Integrative Medicine in Rheumatology:   An Evidence-Based Approach Peter E. Lipsky, MD Chief, Autoimmunity Branch National Institute of Arthritis and Musculoskeletal   and Skin Diseases National Institutes of Health Bethesda, Maryland   Autoimmunity



B. Asher Louden, MD Resident in Dermatology Wake Forest University Baptist Medical Center Winston-Salem, North Carolina   Behçet’s Disease Carlos J. Lozada, MD, FACP, FACR Associate Professor of Medicine University of Miami Miller School of Medicine Director, Rheumatology Fellowship Program   and Rheumatology Clinical Services Jackson Memorial Hospital Miami, Florida   Management of Osteoarthritis Ingrid E. Lundberg, MD, PhD Professor of Medicine and Head, Rheumatology Unit, Department of Medicine Karolinska Institute/Karolinska University Hospital Stockholm, Sweden   Inflammatory Diseases of Muscle and Other Myopathies Reuven Mader, MD Senior Clinical Lecturer B. Rappaport Faculty of Medicine, Technion Israel Institute of Technology Head, Rheumatic Diseases Unit Ha’Emek Medical Center Haifa, Israel   Proliferative Bone Diseases Rashmi M. Maganti, MD Fellow, Division of Rheumatology and Clinical Immunogenetics The University of Texas Health Science Center at Houston Houston, Texas   Rheumatic Manifestations of Human Immunodeficiency Virus Infection Maren Lawson Mahowald, MD Professor of Medicine University of Minnesota Medical School Rheumatology Section Chief Minneapolis VA Medical Center Minneapolis, Minnesota   Management of Chronic Pain



Michael D. Lockshin, MD, MACR Professor of Medicine and Obstetrics-Gynecology Weill Medical College of Cornell University New York, New York   Antiphospholipid Syndrome



Walter P. Maksymowych, MBChB, FRCPC, FACP, FRCP(UK) Professor of Medicine University of Alberta Faculty of Medicine Senior Scientist Alberta Heritage Foundation for Medical Research Edmonton, Alberta, Canada   Ankylosing Spondylitis



Kate R. Lorig, RN, DrPH Professor, Department of Medicine, Division of Immunology and Rheumatology Stanford University School of Medicine Director, Patient Education Research Center Stanford, California   Arthritis Self-Management



Scott David Martin, MD Assistant Professor of Orthopedics Harvard Medical School Attending Staff Physician, Department of Orthopedics Brigham and Women’s Hospital Boston, Massachusetts   Shoulder Pain



CONTRIBUTORS



Helena Marzo-Ortega, MD, MRCP Consultant Rheumatologist and Honorary Senior Lecturer Academic Section of Musculoskeletal Disease Leeds Institute of Molecular Medicine University of Leeds and Chapel Allerton Hospital Leeds, United Kingdom   Undifferentiated Spondyloarthritis and Reactive Arthritis Dennis McGonagle, PhD, FRCPI Professor of Investigative Rheumatology The University of Leeds Leeds, United Kingdom   Undifferentiated Spondyloarthritis and Reactive Arthritis Iain B. McInnes, PhD, FRCP Professor of Experimental Medicine Honorary Consultant Rheumatologist Centre for Rheumatic Diseases, Faculty of Medicine University of Glasgow Glasgow, United Kingdom   Cytokines; Atherosclerosis in Rheumatic Disease; Rheumatoid Factors and Other Autoantibodies in Rheumatoid Arthritis Kevin G. Moder, MD Consultant and Associate Professor Division of Rheumatology and Department of Internal Medicine Mayo Clinic Rochester, Minnesota   History and Physical Examination of the Musculoskeletal System Eamonn S. Molloy, MD, MRCPI Associate Staff Department of Rheumatic and Immunologic Disease Cleveland Clinic Foundation Cleveland, Ohio   Antineutrophil Cytoplasmic Antibody–Associated Vasculitis Kanneboyina Nagaraju, PhD, DVM Associate Professor of Pediatrics George Washington University School of Medicine   and Health Sciences Director, Murine Drug Testing Facility Center for Genetic Medicine Research Children’s Research Institute Children’s National Medical Center Washington, DC   Inflammatory Diseases of Muscle and Other Myopathies Stanley J. Naides, MD Medical Director, Immunology R&D Quest Diagnostics Nichols Institute San Juan Capistrano, California   Viral Arthritis



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Lee S. Newman, MD, MA Professor, Department of Medicine, Division of Allergy and Clinical Immunology and Division of Pulmonary Sciences and Critical Care Medicine Professor of Epidemiology, Department of Preventive Medicine and Biometrics University of Colorado School of Medicine Denver, Colorado   Sarcoidosis Peter A. Nigrovic, MD Instructor in Medicine Harvard Medical School Staff Rheumatologist Department of Rheumatology, Immunology, and Allergy, Brigham & Women’s Hospital Division of Immunology, Children’s Hospital Boston Boston, Massachusetts   Mast Cells Kiran Nistala, MD, MRCP, MSc Clinical Research Fellow in Paediatric Rheumatology Institute of Child Health University College London London, United Kingdome   Juvenile Idiopathic Arthritis James R. O’Dell, MD Larson Professor and Vice-Chairman, Department of Internal Medicine University of Nebraska College of Medicine Chief of Rheumatology and Residency Program Director, Department of Internal Medicine University of Nebraska Medical Center Omaha, Nebraska   Methotrexate, Leflunomide, Sulfasalazine, Hydroxychloroquine, and Combination Therapies Peter R. Oesch, MSc, Dipl PT Head, Department of Ergonomics and Clinical Research Valens Rehabilitation Clinic Valens, Switzerland   Introduction to Physical Medicine and Rehabilitation Yasunori Okada, MD, PhD Professor and Chairman, Department of Pathology Keio University School of Medicine Tokyo, Japan   Proteinases and Matrix Degradation Eugenia C. Pacheco-Pinedo, MD, MSc Postdoctoral Researcher Department of Medicine Molecular Cardiology Research Center Postdoctoral Researcher Department of Physiology Pennsylvania Muscle Institute Philadelphia, Pennsylvania   Muscle: Anatomy, Physiology, and Biochemistry



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CONTRIBUTORS



Richard S. Panush, MD Professor of Medicine Mount Sinai School of Medicine Chair, Department of Medicine Saint Barnabas Medical Center New York, New York   Occupational and Recreational Musculoskeletal   Disorders Thomas Pap, MD Professor of Experimental Medicine Head, Division of Molecular Medicine of Musculoskeletal Tissue University of Münster Institute of Experimental Musculoskeletal Medicine Director University Hospital Münster Münster, Germany   Fibroblasts and Fibroblast-like Synoviocytes Stanford L. Peng, MD, PhD Senior Director, Translational Medicine Leader Clinical Research and Exploratory Development Roche Palo Alto Palo Alto Assistant Clinical Professor, Department of Medicine, Division of Rheumatology–Arthritis University of California, San Francisco, School of Medicine San Francisco, California   Antinuclear Antibodies Harris Perlman, PhD Associate Professor, Department of Molecular Microbiology & Immunology Saint Louis University School of Medicine St. Louis, Missouri   Signal Transduction Jean-Charles Piette, MD Department of Internal Medicine Groupe Hospitalier Pitié-Salpêtrière Paris, France   Relapsing Polychondritis Michael H. Pillinger, MD Chief of Rheumatology New York Hospital for Joint Diseases New York, New York   Neutrophils and Eosinophils Robert S. Pinals, MD Acting Chief, Division of Rheumatology & Connective Tissue Research Professor, Department of Medicine UMDNJ–Robert Wood Johnson Medical School New Brunswick, New Jersey   Felty’s Syndrome



Steven A. Porcelli, MD Weinstock Professor of Microbiology & Immunology   and Professor of Medicine Albert Einstein College of Medicine Bronx, New York   Innate Immunity Mark D. Price, MD, PhD Chief Resident in Orthopedic Surgery Harvard Combined Orthopedic Surgery Program Massachusetts General Hospital Boston, Massachusetts   Foot and Ankle Pain Johannes J. Rasker, MD, PhD Professor Emeritus of Rheumatology Faculty of Behavioral Sciences, Department of Psychology and Communication of Health and Risk University of Twente Enschede, The Netherlands   Fibromyalgia John D. Reveille, MD Professor of Internal Medicine Director, Division of Rheumatology and Clinical Immunogenetics Department of Internal Medicine University of Texas Medical School at Houston Houston, Texas   Rheumatic Manifestations of Human Immunodeficiency Virus Infection W. Neal Roberts, Jr., MD Rheumatology Fellowship Program Director Chas. W. Thomas Professor of Medicine Virginia Commonwealth University School of Medicine, Medical College of Virginia Campus Richmond, Virginia   Psychosocial Management of Rheumatic Diseases James T. Rosenbaum, MD Edward E. Rosenbaum Professor of Inflammation Research and Professor of Ophthalmology, Medicine, and Cell Biology Chair, Division of Arthritis and Rheumatic Diseases Vice-Chair, Department of Ophthalmology Oregon Health & Science University School of Medicine Portland, Oregon   The Eye and Rheumatic Diseases Andrew E. Rosenberg, MD Associate Professor Harvard Medical School Director of Surgical Pathology Massachusetts General Hospital Boston, Massachusetts   Tumors and Tumor-like Lesions of Joints and Related   Structures



CONTRIBUTORS



Clinton T. Rubin, PhD Distinguished Professor and Chair Department of Biomedical Engineering State University of New York Stony Brook University School of Medicine Stony Brook, New York   Biology, Physiology, and Morphology of Bone



David C. Seldin, MD, PhD Professor of Medicine and Microbiology Boston University School of Medicine Director, Amyloid Treatment and Research Program Boston Medical Center Boston, Massachusetts   Amyloidosis



Janet E. Rubin, MD Professor of Medicine and Pharmacology University of North Carolina at Chapel Hill School   of Medicine Chapel Hill, North Carolina   Biology, Physiology, and Morphology of Bone



Jérémie Sellam, MD, PhD Assistant Professor of Rheumatology Paris Universitas – Pierre & Marie Curie Paris VI Assistant Professor of Rheumatology Saint-Antoine Hospital Paris, France   Clinical Features of Osteoarthritis



Holly M. Sackett, MSPH Senior Professional Research Assistant University of Colorado Denver Colorado School of Public Health Denver, Colorado   Sarcoidosis Jane E. Salmon, MD Professor of Medicine Professor of Obstetrics and Gynecology Weill Medical College of Cornell University Attending Physician Hospital for Special Surgery New York Presbyterian Hospital New York, New York   Antiphospholipid Syndrome Jonathan Samuels, MD Instructor in Medicine (Rheumatology) NYU School of Medicine Director, Clinical Immunulogy Laboratory NYU Langone Medical Center   Pathogenesis of Osteoarthritis Naveed Sattar, MBChB, PhD, MRCPath Professor of Metabolic Medicine British Heart Foundation Glasgow Cardiovascular Research Centre University of Glasgow Honorary Consultant Endocrinologist Glasgow Royal Infirmary Glasgow, United Kingdom   Atherosclerosis in Rheumatic Disease John C. Scatizzi, PhD Post-Doctoral Fellow, Division of Rheumatology, Allergy, & Immunology UC San Diego School of Medicine La Jolla, California   Signal Transduction Jose U. Scher, MD Teaching Assistant Department of Medicine, Division of Rheumatology New York University School of Medicine New York, New York   Neutrophils and Eosinophils



xv



John S. Sergent, MD, MACR Professor of Medicine Vice Chair for Education and Residency Program Director Vanderbilt University School of Medicine Nashville, Tennessee   Polyarticular Arthritis; Polyarteritis and Related Disorders; Isolated Angiitis of the Central Nervous System; Arthritis Accompanying Endocrine   and Metabolic Disorders Richard M. Siegel, MD, PhD Investigator, Autoimmunity Branch National Institute of Arthritis and Musculoskeletal   and Skin Diseases National Institutes of Health Bethesda, Maryland   Autoimmunity Karl Sillay, MD Assistant Professor and Director, Functional Surgery, Department of Neurological Surgery University of Wisconsin School of Medicine and Public Health Neurosurgeon, University of Wisconsin Hospital and Clinics, William S. Middleton Memorial Veterans Hospital, St. Mary’s Hospital Medical Center, and Meriter Hospital Madison, Wisconsin   Neck Pain Anna Simon, MD, PhD Clinical Investigator, Department of General Internal Medicine Radboud University Nijmegen Medical Centre Faculty   of Medical Sciences Nijmegen, The Netherlands   Familial Auto-inflammatory Syndromes Dawd S. Siraj, MD, MPH, TM Assistant Professor of Medicine Brody School of Medicine at East Carolina University Director ECU Physicians International Travel Clinic, Section   of Infectious Diseases Greenville, North Carolina   Bacterial Arthritis



xvi



CONTRIBUTORS



Martha Skinner, MD Professor of Medicine Boston University School of Medicine Boston, Massachusetts   Amyloidosis Kathleen A. Sluka, PT, PhD Professor of Physical Therapy Graduate Programs in Physical Therapy and Rehabilitation Science, in Pain Research, and in Neuroscience University of Iowa Carver College of Medicine Iowa City, Iowa   Neurological Regulation of Inflammation C. Michael Stein, MBChB, MRCP Dan May Professor of Medicine and Professor   of Pharmacology Vanderbilt University School of Medicine Nashville, Tennessee   Immunoregulatory Drugs John H. Stone, MD, MPH Clinical Director of Rheumatology Massachusetts General Hospital Boston, Massachusetts   The Classification and Epidemiology of Systemic Vasculitis; Immune Complex–Mediated Small Vessel Vasculitis Bob Sun, MD Instructor, Department of Medicine, Division of Rheumatology Northwestern University Feinberg School of Medicine Attending Rheumatologist Evanston Northwestern Healthcare Evanston, Illinois   Rheumatic Manifestations of Hemoglobinopathies Carrie R. Swigart, MD Assistant Professor, Department of Orthopaedics and Rehabilitation, Hand and Upper Extremity Section Yale University School of Medicine New Haven, Connecticut   Hand and Wrist Pain Zoltán Szekanecz, MD, PhD, DSc Professor, Department of Medicine, Division of Rheumatology and Immunology Institute for Internal Medicine, Rheumatology Division University of Debrecen Medical and Health Science Center Debrecen, Hungary   Cell Recruitment and Angiogenesis Paul P. Tak, MD, PhD Professor of Medicine and Director, Division of Clinical Immunology & Rheumatology Academic Medical Center/University of Amsterdam Faculty of Medicine Amsterdam, The Netherlands   Biologic Markers



Ioannis O. Tassiulas, MD Senior Investigator, Department of Medicine, Division   of Rheumatology University of Crete Medical School Heraklion, Greece   Clinical Features and Treatment of Systemic Lupus Erythematosus H. Guy Taylor, MBChB, MRCP(UK), FRACP, Dipl MSM(Otago) Consultant Rheumatologist Wanganui Hospital Wanganui, New Zealand   Immunoregulatory Drugs Peter C. Taylor, MA, PhD, FRCP Professor of Experimental Rheumatology Head, Clinical Trials Kennedy Institute of Rheumatology Division Faculty of Medicine Imperial College London London, United Kingdom   Cell-Targeted Biologics and Emerging Targets: Rituximab, Abatacept, and Other Biologics Robert Terkeltaub, MD Professor of Medicine Rheumatology Training Program Director and Associate Division Director for Rheumatology–Allergy/ Immunology UC San Diego School of Medicine La Jolla Section Chief, Rheumatology–Allergy VA Medical Center San Diego San Diego, California   Diseases Associated with Articular Deposition of Calcium Pyrophosphate Dihydrate and Basic Calcium Phosphate Crystals Thomas S. Thornhill, MD Professor of Orthopedics Harvard Medical School Chief of Orthopedics Brigham and Women’s Hospital Boston, Massachusetts   Shoulder Pain Helen Tighe, BSc, PhD Associate Adjunct Professor, Department of Medicine, Division of Rheumatology, Allergy, and Immunology UC San Diego School of Medicine La Jolla, California   Rheumatoid Factors and Other Autoantibodies in Rheumatoid Arthritis Betty P. Tsao, PhD Professor of Medicine, Department of Medicine, Division of Rheumatology David Geffen School of Medicine at UCLA Los Angeles, California   Pathogenesis of Systemic Lupus Erythematosus



CONTRIBUTORS



Peter Tugwell, MSc, MD, FRCPC Professor of Medicine, Department of Medicine Ottawa Health Research Institute University of Ottawa Faculty of Medicine Ottawa, Ontario, Canada   Assessment of Health Outcomes Zuhre Tutuncu, MD Associate Professor, Department of Rheumatology, Allergy, and Immunology UC San Diego School of Medicine La Jolla, California   Anticytokine Therapies Katherine S. Upchurch, MD Associate Professor of Medicine University of Massachusetts Medical School Clinical Chief, Division of Rheumatology UMass Memorial Medical Center Worcester, Massachusetts   Hemophilic Arthropathy Wim B. Van den Berg, PhD Professor of Experimental Rheumatology, Rheumatology Research, and Advanced Therapeutics Radboud University Nijmegen Medical Centre Faculty of Medical Sciences Nijmegen, The Netherlands   Animal Models of Inflammatory Arthritis



John Varga, MD Hughes Distinguished Professor of Medicine Department of Medicine, Division of Rheumatology Northwestern University Feinberg School of Medicine Chicago, Illinois   Systemic Sclerosis and the Scleroderma-Spectrum Disorders Philippe Vinceneux, MD Medecine Interne 2 Hospital Pitié-Salpêtrière Paris, France   Relapsing Polychondritis Benjamin W.E. Wang, MD, FRCPC Associate Professor, Department of Medicine, Division   of Rheumatology University of Tennessee College of Medicine Memphis, Tennessee   Nonsteroidal Anti-inflammatory Drugs Lucy R. Wedderburn, MD Reader in Paediatric Rheumatology Rheumatology Unit, Institute of Child Health–University College London London, United Kingdom   Juvenile Idiopathic Arthritis



Filip Van den Bosch, MD, PhD Rheumatologist University Hospital Gent—Department of Rheumatology Ghent, Belgium   Undifferentiated Spondyloarthritis and Reactive Arthritis



Barbara N. Weissman, MD Professor of Radiology Harvard Medical School Director, Radiology Residency Program Vice Chair, Department of Radiology Brigham & Women’s Hospital Boston, Massachusetts   Imaging Modalities in Rheumatic Disease



Désirée M. F. M. Van der Heijde, MD, PhD Professor of Rheumatology, Department of   Rheumatology Leiden University Faculty of Medicine Leiden, The Netherlands   Clinical Trial Design and Analysis; Ankylosing   Spondylitis



Victoria P. Werth, MD Professor of Dermatology and Medicine University of Pennsylvania School of Medicine Chief of Dermatology Philadelphia VA Medical Center Philadelphia, Pennsylvania   The Skin and Rheumatic Diseases



Sjef M. van der Linden, MD Professor of Rheumatology, Department of Medicine University of Maastricht Faculty of Health, Medicine,   and Life Sciences CAPHRI Research Institute Head, Division of Rheumatology, Department of   Medicine University Hospital of Maastricht Maastricht, The Netherlands   Ankylosing Spondylitis



Karin N. Westlund-High, PhD Professor, Department of Physiology University of Kentucky College of Medicine Lexington, Kentucky   Neurological Regulation of Inflammation



Jos W.M. van der Meer, MD, PhD, FRCP Department of General Internal Medicine Radboud University Nijmegen Medical Centre Nijmegen, The Netherlands   Familial Auto-inflammatory Syndromes



xvii



Michael S. Wildstein, MD President Wildstein Spine Center Charleston, South Carolina   Low Back Pain



xviii



CONTRIBUTORS



Christopher M. Wise, MD W. Robert Irby Professor of Medicine Department of Medicine, Division of Rheumatology, Allergy, and Immunology Virginia Commonwealth University School of Medicine, Medical College of Virginia Campus Richmond, Virginia   Arthrocentesis and Injection of Joints and Soft Tissue Frederick Wolfe, MD Clinical Professor of Medicine University of Kansas School of Medicine Director, National Data Bank for Rheumatic   Diseases Wichita, Kansas   Fibromyalgia Frank A. Wollheim, MD, PhD, FRCP Emeritus Professor, Department of Rheumatology Lund University Faculty of Medicine/Lund University Hospital Lund, Sweden   Enteropathic Arthritis Patricia Woo, MBBS, BSc, PhD, CBE, FRCP, FMedSci Professor of Paediatric Rheumatology Faculty of Medicine University College London London, United Kingdom   Juvenile Idiopathic Arthritis Anthony D. Woolf, BSc, MBBS, FRCP Professor of Rheumatology Peninsula College of Medicine and Dentistry, Universities of Exeter & Plymouth Exeter Consultant Rheumatologist Duke of Cornwall Rheumatology Unit, Royal Cornwall Hospital Truro, United Kingdom   Economic Burden of Rheumatic Diseases



Robert L. Wortmann, MD Professor of Medicine Dartmouth-Hitchcock Medical Center Lebanon, New Hampshire   Gout and Hyperuricemia David Tak Yan Yu, MD Professor of Medicine David Geffen School of Medicine at UCLA Los Angeles, California   Undifferentiated Spondyloarthritis and Reactive Arthritis John B. Zabriskie, MD Professor Emeritus Rockefeller University New York, New York   Poststreptoccocal Arthritis and Rheumatic Fever Robert B. Zurier, MD Professor, Department of Medicine, Division   of Rheumatology University of Massachusetts Medical School Worcester, Massachusetts   Prostaglandins, Leukotrienes, and Related Compounds Anne-Marie Zuurmond, PhD Biosciences Division TNO Quality of Life Leiden, The Netherlands   Biologic Markers



PREFACE



“Plus ça change, plus c’est la même chose” –Jean-Baptiste Alphonse Karr As we, the editors, worked on the 8th edition of Kelley’s Textbook of Rheumatology, we were struck by Monsieur Karr’s quote from 160 years ago. This textbook continues to change and evolve. The incomparable Ted Harris, who was editor-in-chief of the 7th edition, has stepped down, leaving Gary Firestein to fill his shoes. For the first time, a European editor, Iain McInnes, joined the group and helped create a truly international edition. Full color was introduced, new chapter formats were designed to assure a consistent look and feel for each topic, and algorithms for diagnosis and treatment as well as key point boxes were included. In addition, new authors were added to the list of luminaries that already contribute to the book, and there was a major effort to provide greater availability through electronic versions of reference material and on-line access. While change has been in the air regarding many aspects of the book, some things never vary. The book was initially designed decades ago to provide scholarly rigor to the field of rheumatology and to offer definitive reviews of scientific advances as they apply to clinical medicine. This remained the touchstone of our enterprise for the past 4 years, just as it was for the previous seven editions. The painstaking job of identifying the premier authors to present definitive







i­nformation on each topic required months of work, culminating in an editors’ meeting in Costa Rica to finalize the chapters (well, it wasn’t all work!). The arduous process of guiding, reviewing, and editing the outstanding contributions of our authors was time consuming but paled compared with the efforts that the authors put into creating their chapters. The thread connecting the past to the present was also evident in the continued effort of our valued co-editors John Sergent, Ralph Budd, and Shaun Ruddy. Our trusted colleagues at Elsevier, including Cathy Carroll and Kimberly Murphy, were always available and suffered with us in Costa Rica as well. The present looks very encouraging indeed. Kelley’s Textbook of Rheumatology, 8th edition, is a beautiful tome that is designed to carry on the tradition of being the definitive rheumatology resource. Looking to the future, we expect that this work will continue to evolve and change. New editors, new science, new authors, and new technology will be the rule rather than the exception. As you begin to use this edition, please know that it is truly a labor of love. We have enjoyed the experience and hope that it is as valuable to you as the previous editions. The Editors







xix



Part



1



Structure and Function of Bone, Joints, and Connective Tissue



1



Biology of the Normal Joint Steven R. Goldring  •  Mary B. Goldring



KEY POINTS



CLASSIFICATION OF JOINTS



Condensation of mesenchymal cells, which differentiate into chondrocytes, results in formation of the cartilage anlagen, which provides the template for the developing skeleton.



Human joints provide the structures by which bones join with one another and may be classified according to the histologic features of the union and the range of joint motion. There are three classes of joint design: (1) synovial or ­diarthrodial joints (Fig. 1-1), which articulate with free movement, have a synovial membrane lining the joint cavity, and contain synovial fluid; (2) amphiarthroses, in which adjacent bones are separated by articular cartilage or a fibrocartilage disk and are bound by firm ligaments permitting limited motion (e.g., pubic symphysis, intervertebral disks of vertebral bodies, distal tibiofibular articulation, and ­sacroiliac joint articulation with pelvic bones); and (3) synarthroses, which are found only in the skull (suture lines) where thin, fibrous tissue separates adjoining cranial plates that interlock to prevent detectable motion before the end of normal growth, yet permit growth in childhood and adolescence.1 Joints also can be classified according to the connective tissues present. Symphyses have a fibrocartilaginous disk separating bone ends that are joined by firm ligaments (e.g., symphysis pubis and intervertebral joints). In synchondroses, the bone ends are covered with articular cartilage, but there is no synovium or significant joint cavity (e.g., sternomanubrial joint). In syndesmoses, the bones are joined directly by fibrous ligaments without a cartilaginous interface (the distal tibiofibular articulation is the only joint of this type outside the cranial vault). In synostoses, bone bridges are formed between bones, producing ankylosis. The synovial joints are classified further according to their shapes, which include ball-and-socket (hip), hinge (interphalangeal), saddle (first carpometacarpal), and plane (patellofemoral) joints. These configurations reflect the varying functions, as the shapes and sizes of the opposing surfaces determine the direction and extent of motion. The various designs permit flexion, extension, abduction, adduction, or rotation. Certain joints can act in one (humeroulnar), two (wrist), or three (shoulder) axes of motion. This chapter concentrates on the developmental biology and relationship between structure and function of a “prototypic,” “normal” human diarthrodial joint—the joint most likely to develop arthritis. Most research that has been done



During development of the synovial joint, growth differentiation factor-5 regulates interzone formation, and interference with movement of the embryo during development impairs joint cavitation. Members of the bone morphogenetic protein/transforming growth factor-β, fibroblast growth factor, and Wnt families and the parathyroid hormone–related peptide/Indian hedgehog axis are essential for joint development and growth plate formation. The synovial lining of diarthrodial joints is a thin layer of cells lacking a basement membrane and consisting of two principal cell types—macrophages and fibroblasts. The articular cartilage receives its nutritional requirements via diffusion from the synovial fluid, and interaction of the cartilage with components of the synovial fluid contributes to the unique low-friction surface properties of the articular cartilage.



The normal joint is a specialized, integrated structure consisting of multiple connective tissue elements, including muscles, tendons, ligaments, synovium and capsule, cartilage, and bone, organized in a manner that permits stability and movement of the human skeleton. The joint structures are positioned to distribute normal mechanical stresses optimally and are organized for low-friction load bearing. Deviations from normal structure and physiology of joint tissues have been implicated in the pathogenesis of various forms of arthritis. The differentiation of articular tissues during embryonic development dictates their capacity to respond to insults in later life. Physiologic cellular processes involved in normal joint development, such as differentiation, angiogenesis, macrophage recruitment, and fibroblast proliferation, may reappear in the mature joint and contribute to the pathogenesis of joint disease. Knowledge of the development, structure, and function of normal joint tissues is essential for understanding the underlying mechanisms involved in the pathogenesis of human joint diseases.











GOLDRING 



| 



Biology of the Normal Joint



Cartilage Bone



Homogeneous 3-Layered interzone Mesenchyme Perichondrium Synovial interzone Blastema mesenchyme Cartilage



Tide mark



Periosteum



A



B



D Cavities Capsule Synovium Figure 1-1  A normal human interphalangeal joint, in sagittal section, as an example of a synovial, or diarthrodial, joint. The tidemark represents the calcified cartilage that bonds articular cartilage to the subchondral bone plate. (From Sokoloff L, Bland JH: The Musculoskeletal System. Baltimore, Williams & Wilkins, 1975. © 1975, the Williams & Wilkins Co, Baltimore.)



concerns the knee because of its accessibility, but other joints are described when appropriate.



DEVELOPMENTAL BIOLOGY OF THE DIARTHRODIAL JOINT Skeletal development is initiated by the differentiation of mesenchymal cells that arise from three sources: (1) neural crest cells of the neural ectoderm that gives rise to craniofacial bones; (2) the sclerotome of the paraxial mesoderm, or somite compartment, which forms the axial skeleton; and (3) the somatopleure of the lateral plate mesoderm, which yields the skeleton of the limbs.2 The appendicular skeleton develops in the human embryo from limb buds, which are first visible at around 4 weeks of gestation. Structures resembling adult joints are generated at approximately 4 to 7 weeks of gestation.3 Many other crucial phases of musculoskeletal development follow, including vascularization of epiphyseal cartilage (8 to 12 weeks), appearance of villous folds in synovium (10 to 12 weeks), evolution of bursae (3 to 4 months), and appearance of periarticular fat pads (4 to 5 months). The upper limbs develop approximately 24 hours earlier than the analogous portions of the lower limbs. Proximal structures, such as the glenohumeral joint, develop before more distal ones, such as the wrist and hand. As a consequence, insults to embryonic development during limb formation affect a more distal portion of the upper limb than of the lower limb. Long bones form as a result of replacement of the cartilage template by endochondral ossification. The stages of limb development are well described by O’Rahilly and Gardner3,4 and are shown in Figure 1-2. The ­developmental sequence of the events occurring during



C



E Articular capsule



Articular cavity



Synovial tissue and fold



Figure 1-2  The development of a synovial joint. A, Condensation. Joints develop from the blastema, not the surrounding mesenchyme.  B, Chondrification and formation of the interzone. The interzone remains avascular and highly cellular. C, Formation of synovial mesenchyme. Synovial mesenchyme forms from the periphery of the interzone and is invaded by blood vessels. D, Cavitation. Cavities are formed in the central and peripheral interzone and merge to form the joint cavity. E, The  mature joint. (From O’Rahilly R, Gardner E: The embryology of movable joints. In Sokoloff L (ed): The Joints and Synovial Fluid, Vol 1. New York, ­Academic Press, 1978.)



synovial joint formation and some of the regulatory factors and extracellular matrix components involved are summarized in Figures 1-3 and 1-4. INTERZONE FORMATION AND JOINT CAVITATION The morphology of the developing synovial joint and the process of joint cavitation have been described in many classic studies done on the limbs of mammalian and avian embryos.5 In the human embryo, cartilage condensations, or chondrifications, can be detected at stage 17, when the embryo is small, approximately 11.7 mm long.3,4 In the region of the future joint, following formation of the homogeneous chondrogenic interzone at 6 weeks (stages 18 and 19), a three-layered interzone is formed at approximately 7 weeks (stage 21), which consists of two chondrogenic, perichondrium-like layers that cover the opposing surfaces of the cartilage anlagen and are separated by a narrow band of densely packed cellular blastema that remains and forms the interzone. Cavitation begins in the central interzone at about 8 weeks (stage 23). Although these cellular events associated with joint formation have been recognized for many years, only more recently have the genes regulating these processes been elucidated. These genes include growth differentiation factor (GDF)-5, Wnt-14, bone morphogenetic protein (BMP)-2, BMP-4, BMP-6, BMP-7, and the GDF-BMP antagonists.5-8 In addition, joint formation is ­accompanied by the expression of several fibroblast growth factor (FGF) family members, including FGF-2 and FGF-4.9 The balance of signaling between BMP and FGF determines the rate of



PART 1 



Mesenchymal cell condensation



| 



STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



Chondrocyte proliferation



Chondrocyte differentiation



TGF-β Wnt-3A, 7A FGF-2, 4, 8,10 Sonic Hh BMP-2, 4, 7



IGF-1 FGF-2/FGFR2 BMP-2, 4, 7, 14



HoxA, HoxD Sox9 Gli3



PTHrP



FGFR2



FGF-2



FGFR3



Ihh BMP-2



Ossification



FGF-18/FGFR3 BMP-2, 7 Bone PTHrP collar Ihh/Ptc



VEGF FGF-2/FGF-R1 Wnt14/ β-catenin



Stat1 Gli3, 2 Runx2 Fra2/JunD



Runx2 Osterix TCF/Lef1



Epiphyseal ossification center (secondary) Diaphyseal ossification center (primary)



Growth plate



Periarticular (resting) Proliferating



BMP-7



FGF-18 Ptc



Perichondrium



FGF-2 BMPs



Gli



Sox9, 5, 6



Chondrocyte hypertrophy and vascular invasion







Prehypertrophic



FGFR1 Hypertrophic



BMP-6



Collagen II, IX, XI Aggrecan COMP Collagen X Osteocalcin



Figure 1-3  The stages of diarthrodial joint formation, and the temporal pattern of expression of the genes involved in regulation at different stages. Subperiosteal ring



TGF-β FGF-2,4,8,10 Wnt3A,7A Shh BMP-2,4,7 Gli3 HoxA, D r-Fng Lmx1b RA Mesenchymal condensation



Sox9,5,6 IGF-1 FGF-2,18 BMP-2,4,7,14 PTHrP Ihh



Diaphyseal ossification center



Wnt14 GDF-5 BMP-2,4 FGF-2 Runx2 Cux1 Erg5



Epiphyseal ossification center Synovial capsule Hyaluronan CD44



Interzone formation Interzone Joint initiation and chondrocyte formation and ossification differentiation Figure 1-4  Development of long bones from cartilage anlagen.



proliferation, adjusting the pace of the differentiation.10 Two transcription factors, Cux-1, a homeobox factor, and the ETS factor ERG/C-1-1, are expressed concurrently with GDF-5 and Wnt-14 at the onset of joint formation.11,12 Hartmann and Tabin13 have proposed two major roles for Wnt-14. First, it acts at the onset of joint formation as a negative regulator of chondrogenesis. Second, it facilitates interzone formation and cavitation by inducing the expression of GDF-5 (also known as cartilage-derived morphogenetic protein-1 [CDMP-1]), Wnt-4, chordin, and the hyaluronan receptor, CD44.13-15 Paradoxically, application of GDF-5 to developing joints in mouse embryo limbs in organ culture causes joint fusion,16 suggesting that temporospatial interactions among distinct cell populations are important for the correct response. The current view



Cavitation



C-1-1 Articular cartilage



Joint maturation



is that GDF-5 is required at the early stages of condensations, where it stimulates recruitment and differentiation of chondrogenic cells, and later, when its expression is restricted to the interzone. The distribution of collagen types and keratan sulfate in developing avian and rodent joints has been characterized by immunohistochemistry.17-21 Collagen types I and III characterize the matrix produced by mesenchymal cells, which switch to the production of types II, IX, and XI collagens that typify the cartilaginous matrix at the time of condensation.22 The messenger RNAs encoding the small proteoglycans, biglycan and decorin, may be expressed at this time, but the proteins do not appear until after cavi­ tation in the regions destined to become articular cartilage.23 The interzone regions are marked by the expression







GOLDRING 



| 



Biology of the Normal Joint



C



A



B



C C



C



D



C C



E



F



Figure 1-5  In situ hybridization of a 13-day-old (stage 39) chicken embryo middle digit, proximal interphalangeal joint, midfrontal sections. A, Brightfield image showing developing joint and capsule (C). B, Equivalent paraffin section of opposite limb of same animal, showing onset of cavitation laterally (arrow). C, Expression of type IIA collagen mRNA in articular surface cells, perichondrium, and capsule. D, Type IIB collagen mRNA is expressed only in chondrocytes of the anlagen. E, Type XI collagen mRNA is expressed in the surface cells, perichondrium, and capsule, with lower levels in chondrocytes. F, Type I collagen mRNA is present in cells of the interzone and capsule. C through F images are dark field. Calibration bar = 1 μm. (From Nalin AM, Greenlee TK Jr, Sandell LJ: Collagen gene expression during development of avian synovial joints: Transient expression of types II and XI collagen genes in the joint capsule. Develop Dyn 203:352-362, 1995.)



of type IIA ­ collagen by chondrocyte progenitors in the perichondrial layers, type IIB and XI collagens by overt­ chondrocytes in the cartilage anlagen, and type I collagen in the interzone and in the developing capsule and perichondrium (Fig. 1-5).24 The interzone region contains cells in two outer layers that are destined to differentiate into chondrocytes and become incorporated into the epiphyses, and in a thin intermediate zone that are programmed to undergo joint cavitation and may remain as articular chondrocytes.25 Fluid and macromolecules accumulate in this space and create a nascent synovial cavity. Blood vessels appear in the surrounding capsulosynovial blastemal mesenchyme before separation of the adjacent articulating surfaces.26 Although it was first assumed that these interzone cells should undergo necrosis or programmed cell death (apoptosis),27 some investigators have found no evidence of DNA ­fragmentation preceding



cavitation.24,25,28,29 There also is no evidence that metalloproteinases are involved in loss of tissue strength in the region undergoing cavitation.30 Instead, the actual joint cavity seems to be formed by mechanospatial changes induced by the synthesis of hyaluronan via uridine diphosphoglucose dehydrogenase (UDPGD) and hyaluronan synthase. Interaction of hyaluronan with its cell surface receptor, CD44, modulates cell migration, but it is thought that the accumulation of hyaluronan and the associated mechanical influences play the major role in forcing the cells apart and inducing rupture of the intervening extracellular matrix by tensile forces.20,31 This mechanism accounts for the observation that joint cavitation is incomplete in the absence of movement.32,33 Equivalent data from human embryonic joints are difficult to obtain. In all large joints in humans, complete joint cavities are apparent at the beginning of the fetal period.



PART 1 



| 



STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



CARTILAGE FORMATION AND ENDOCHONDRAL OSSIFICATION The skeleton develops from the primitive, avascular, densely packed cellular mesenchyme, termed the skeletal blastema. Common precursor mesenchymal cells divide into chondrogenic, myogenic, and osteogenic lineages that determine the differentiation of cartilage centrally, muscle peripherally, and bone. The surrounding tissues, particularly epithelium, influence the differentiation of mesenchymal progenitor cells to chondrocytes in cartilage anlagen. The cartilaginous nodules appear in the middle of the blastema, and simultaneously cells at the periphery become flattened and elongated to form the perichondrium. In the vertebral column, cartilage disks arise from portions of the somites surrounding the notochord, and nasal and auricular cartilage and the embryonic epiphysis form from the perichondrium. In the limb, the cartilage remains as a resting zone that later becomes the articular cartilage, or undergoes terminal hypertrophic differentiation to become calcified (growth plate formation) and is replaced by bone (endochondral ossification). The latter process requires extracellular matrix remodeling and vascularization (angiogenesis). These events are controlled exquisitely by cellular interactions with the surrounding matrix, growth and differentiation factors, and other environmental factors that initiate or suppress cellular signaling pathways and transcription of specific genes in a temporospatial manner. Condensation and Limb-Bud Formation Formation of the cartilage anlage occurs in four stages: (1) cell migration, (2) aggregation regulated by mesenchymal-epithelial cell interactions, (3) condensation, and (4) overt chondrocyte differentiation, or chondrification.3,4,34 Interactions with the epithelium determine mesenchymal cell recruitment and migration, proliferation, and condensation.34-36 The aggregation of chondroprogenitor mesenchymal cells into precartilage condensations was first described by Fell37 and depends on signals initiated by cell-cell and cell-matrix interactions, the formation of gap junctions, and changes in the cytoskeletal architecture. Before condensation, the prechondrocytic mesenchymal cells produce extracellular matrix that is rich in hyaluronan and type I collagen and type IIA collagen, which contains the exon2-encoded aminopropeptide found in noncartilage collagens.38 The initiation of condensation is associated with increased hyaluronidase activity and the appearance of the cell adhesion molecules, neural cadherin (N-cadherin) and neural cell adhesion molecule (N-CAM), which facilitate cell-cell interactions. Before chondrocyte differentiation, the cell-matrix interactions are facilitated by fibronectin binding to syndecan, downregulating N-CAM and setting the condensation boundaries. Increased cell proliferation and extracellular matrix remodeling, with the disappearance of type I collagen, fibronectin, and N-cadherin, and the appearance of tenascins, matrilins, and thrombospondins, including cartilage oligomeric protein, initiate the transition from chondroprogenitor cells to a fully committed chondrocyte.2,36,39,40 N-cadherin and N-CAM disappear in differentiating chondrocytes and are detectable later only in perichondrial cells.







The differentiated chondrocytes can proliferate and undergo the complex process of hypertrophic maturation or remain within cartilage elements in articular joints. Zwilling41 proposed that positional information for organization of the limb bud was imparted by diffusible agents generated at the tip of the limb bud and along its posterior margin, promoting the development of a cartilaginous anlage along proximal-distal and anterior-posterior axes. Limb buds develop from the lateral plate mesoderm.42 The patterning of limb mesenchyme is due to interactions between the mesenchyme and the overlying epithelium.35 The embryonic limb possesses two signaling centers, the apical ectodermal ridge (AER) and the zone of polarizing activity (ZPA), which produce signals responsible for directing the proximal-distal outgrowth (AER)and anterior-­posterior patterning (ZPA).2,39 Much of the current understanding of limb development is based on early studies in chickens and more recently in mice. The regulatory events are controlled by interacting patterning systems involving FGF, hedgehog, BMP, and Wnt pathways, each of which functions sequentially over time (see Fig. 1-3).42 Wnt signaling via β-catenin is required to induce FGFs, such as FGF-10 and FGF-8, which act in positive feedback loops.42,43 FGF-2, FGF-4, and FGF-8 (induced by Wnt-3A44), from the specialized epithelial cells in the AER that are covering the limb-bud tip, control proximaldistal (shoulder/finger) outgrowth.45 The homeobox (Hox) transcription factors encoded by the HoxA and HoxD gene clusters, which are crucial for the early events of limb patterning in the undifferentiated mesenchyme, are required for the expression of FGF-8 and Sonic hedgehog (Shh),46 and they modulate the proliferation of cells within the condensations.34 Among the Hox genes, Hoxa13 and Hoxd13 enhance and Hoxa11 and Hoxd11 suppress early events in the formation of the cartilage anlagen. Wnt7a is expressed early during limb bud development where it acts to maintain Shh expression.42 Shh, produced by a small group of cells in the posterior zone of the ZPA (in response to retinoic acid in the mesoderm47 and FGF-4 in the AER48), plays a key role in directing anterior-posterior (e.g., little finger/thumb) patterning47,49 and stimulating expression of BMP-2, BMP-4, BMP-7, and Hox genes.50-52 Shh signaling, which is required for early limb patterning, but not for limb formation, is mediated by the Shh receptor Patched (Ptc1), which activates another transmembrane protein, Smoothened (Smo), and inhibits processing of the Gli3 transcription factor to a transcriptional repressor.43,53 Dorsal-ventral (e.g., knuckles/palm) patterning depends on the secretion of Wnt-7A54 and expression of the following transcription factors: radical fringe (r-Fng) by the dorsal ectoderm, and engrailed (En-1) and Lmx1b (which is induced by Wnt-7A) by the ventral endoderm.43,55 BMP-2, BMP-4, and BMP-7 coordinately regulate the patterning of limb elements within the condensations depending on the temporal and spatial expression of BMP receptors, involving SMAD-dependent and SMAD­independent signaling and BMP antagonists, such as noggin and chordin.42,56-58 In vitro and in vivo studies have shown that BMP signaling is required for the formation of precartilaginous condensations and for the differentiation of precursors into chondrocytes.59 Growth of the condensation ceases when noggin inhibits BMP signaling and permits







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overt ­differentiation to chondrocytes, which are often designated as “chondroblasts.” The cartilage formed serves as a template for formation of cartilage elements in the vertebra, sternum, and rib, and for limb elongation or endochondral bone formation. Molecular Signals in Cartilage Morphogenesis and Growth Plate Development The cartilage anlagen grow by cell division and deposition of the extracellular matrix and by apposition of proliferating cells from the inner chondrogenic layer of the perichondrium. The nuclear transcription factor, Sox9, is one of earliest markers expressed in cells undergoing condensation and is required for the subsequent stage of chondrogenesis characterized by the deposition of matrix containing collagens II, IX, and XI and aggrecan in the cartilage anlagen.60,61 Two additional Sox family members, L-Sox5 and Sox6, which are not present in early mesenchymal condensations, but are coexpressed with Sox9 during chondrocyte differentiation,62 have a high degree of sequence identity with each other, but have no sequence homology with Sox9 except in the HMG box. They can form homodimers or heterodimers, which bind more efficiently to pairs of HMG box sites than to single sites, and in contrast to Sox9, they contain no transcriptional activation domain. The expression of SOX proteins depends on BMP signaling via BMPR1A and BMPR1B, which are functionally redundant and active in chondrocyte condensations, but not in the perichondrium.59 L-Sox5 and Sox6 are required for the expression of Col9a1, aggrecan, link protein, and Col2a1 during overt chondrocyte differentiation.63 The runt-domain transcription factor, Runx2 (also known as core binding factor, Cbfa1), also is expressed in all condensations, including those that are destined to form bone.64-66 Throughout chondrogenesis, the balance of signaling by BMPs and FGFs determines the rate of proliferation, adjusting the pace of the differentiation.10 In the long bones, long after condensation, BMP-2, BMP-3, BMP-4, BMP-5, and BMP-7 are expressed primarily in the perichondrium, and only BMP-7 is expressed in the proliferating chondrocytes.10 BMP-6 is found later exclusively in hypertrophic chondrocytes along with BMP-2. More than 23 FGFs have been identified so far.67 The specific ligands that activate each FGF receptor (R) during chondrogenesis in vivo have been difficult to identify because the signaling depends on the temporal and spatial location of not only the ligands, but also the receptors.68 FGFR2 is upregulated early in condensing mesenchyme and is present later in the periphery of the condensation along with FGFR1, which is expressed in surrounding loose mesenchyme. FGFR3 is associated with proliferation of chondrocytes in the central core of the mesenchymal condensation and may overlap with FGFR2. Proliferation of chondrocytes in the embryonic and postnatal growth plate is regulated by multiple mitogenic stimuli, including FGFs, which converge on the cyclin D1 gene.69 In the growth plate, FGFR3 serves as a master inhibitor of chondrocyte proliferation via phosphorylation of the Stat1 transcription factor, which increases the expression of the cell cycle inhibitor p21.70 More recent studies suggest that FGF-18 is the preferred ligand of FGFR3 because Fgf18-deficient mice have an expanded zone of



proliferating chondrocytes similar to that in Fgfr3-deficient mice, and that FGF-18 can inhibit Indian hedgehog (Ihh) expression.71 FGF18 and FGF9 are expressed in the perichondrium and periosteum and form a functional gradient from the proximal proliferating zone, where FGF18 acts via FGFR3 to downregulate proliferation and subsequent maturation.71,72 FGF18 and FGF9 interact with FGFR1 in the prehypertrophic and hypertrophic zones, where more recent evidence indicates that they regulate vascular invasion by inducing the expression of vascular endothelial growth factor (VEGF) and VEGFR1. As the epiphyseal growth plate develops, FGFR3 disappears, and FGFR1 expression is upregulated in prehypertrophic and hypertrophic chondrocytes, suggesting a role for FGFR1 in the regulation of cell survival and differentiation and possibly cell death.68 The proliferation of chondrocytes in the lower proliferative and the prehypertrophic zones is under the control of a local negative feedback loop involving signaling by parathyroid hormone related protein (PTHrP) and Ihh.73 Ihh expression is restricted to the prehypertrophic zone, and the PTHrP receptor is expressed in the distal zone of periarticular chondrocytes. The adjacent, surrounding perichondrial cells express the Hedgehog receptor patched (ptc), which on Ihh binding, similar to Shh in the mesenchymal condensations, activates Smo and induces Gli transcription factors, which can feedback regulate Ihh target genes in a positive (Gli1 and Gli2) or negative (Gli3) manner.74 Ihh induces expression of PTHrP in the perichondrium,75 and PTHrP signaling stimulates cell proliferation via its receptor expressed in the periarticular chondrocytes.76 These interactions are modulated by a balance of BMP and FGF signaling that adjusts the pace of chondrocyte terminal differentiation to the proliferation rate.10 FGF-18 or FGFR3 signaling can inhibit Ihh expression,71 and BMP signaling upregulates the expression of Ihh in cells that are beyond the range of the PTHrP-induced signal.10 Evidence indicates that Ihh acts independently of PTHrP on periarticular chondrocytes to stimulate differentiation of columnar chondrocytes in the proliferative zone, whereas PTHrP acts by preventing premature differentiation into prehypertrophic and hypertrophic chondrocytes, suppressing premature expression of Ihh.77 Ihh and PTHrP, by transiently inducing proliferation markers and repressing differentiation markers, function in a temporospatial manner to determine the number of cells that remain in the chondrogenic lineage versus the number that enter the endochondral ossification pathway.73,78 Endochondral Ossification The development of long bones from the cartilage anlagen occurs by a process termed endochondral ossification, which involves terminal differentiation of chondrocytes to the hypertrophic phenotype, cartilage matrix calcification, vascular invasion, and ossification (see Fig. 1-4).28,78-80 This process is initiated when the cells in the central region of the anlage begin to hypertrophy, increasing cellular fluid volume by almost 20 times. Ihh plays a pivotal role in regulating endochondral bone formation by synchronizing perichondrial maturation with chondrocyte hypertrophy, which is essential for initiating the process of vascular invasion.81 Ihh is expressed in prehypertrophic chondrocytes as they exit the proliferative phase and enter the hypertrophic



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STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



phase, at which time they begin to express the hypertrophic chondrocyte marker, type X collagen (Col10a1) and alkaline phosphatase. These cells are responsible for laying down the cartilage matrix that subsequently undergoes mineralization. Runx2, which serves as a positive regulatory factor in chondrocyte maturation to hypertrophy,82 is expressed in the adjacent perichondrium and in prehypertrophic chondrocytes, but less in late hypertrophic chondrocytes,83,84 overlapping with Ihh, Col10a1, and BMP-6.78,85 BMP-induced Smad1 interacts with Runx2, and Runx2 and Smad1 are important for chondrocyte hypertrophy.82,86,87 An essential role for Runx2 in the process of chondrocyte hypertrophy is supported by the observation that the late stages of chondrocyte hypertrophy are blocked in Runx2-deficient mice.65,88 Interactions with components of the extracellular matrix also contribute to regulation of the process of chondrocyte hypertrophy. Matrix metalloproteinase (MMP)-13, a downstream target of Runx2, is expressed by terminal hypertrophic chondrocytes,89-92 and MMP-13 deficiency results in significant interstitial collagen accumulation leading to the delay of endochondral ossification in the growth plate with increased length of the hypertrophic zone.93,94 In contrast, Col10a1 knockout mice and transgenic mice with a dominant interference Col10a1 mutation have subtle growth plate phenotypes with compressed proliferative and hypertrophic zones and altered mineral deposition.95 Mutations in the COL10A1 gene are associated with the dwarfism observed in human chondrodysplasias. These mutations affect regions of the growth plate that are under great mechanical stress, and it has been suggested that the defect in skeletal growth may be due partly to alteration of the mechanical integrity of the pericellular matrix in the hypertrophic zone, although a role for defective vascularization also has been proposed.96 The extracellular matrix remodeling that accompanies chondrocyte terminal differentiation is thought to induce an alteration in the environmental stress experienced by hypertrophic chondrocytes, which eventually undergo apoptosis.78,97,98 Together these studies indicate that the composition and remodeling of the extracellular matrix play an important role in processes associated with chondrocyte hypertrophy, vascular invasion, and, as discussed subsequently, osteoblast recruitment and subsequent bone formation.91 Vascular invasion of the hypertrophic zone is required for the replacement of calcified cartilage by bone.85,92 The angiogenic factor, VEGF, promotes vascular invasion by specifically activating localized receptors, including Flk expressed in endothelial cells in the perichondrium or surrounding soft tissues, neuropilin 1 (Npn1) expressed in late hypertrophic chondrocytes, or Npn2 expressed exclusively in the perichondrium.28 VEGF is expressed as three different isoforms: VEGF188, a matrix-bound form, is essential for metaphyseal vascularization, whereas the soluble form, VEGF120 (VEGFA), regulates chondrocyte survival and epiphyseal cartilage angiogenesis.99-101 VEGF164 can be either soluble or matrix bound and may act directly on chondrocytes via Npn2. VEGF is released from the extracellular matrix by MMPs, including MMP-9, membranetype (MT)1-MMP (MMP-14), and MMP-13. MMP-9 is expressed by endothelial cells that migrate into the central region of the hypertrophic cartilage.91 MMP-14, which has a broader range of expression than MMP-9, is essential for







chondrocyte proliferation and secondary ossification,102 whereas MMP-13 is found exclusively in late hypertrophic chondrocytes.83 These events of cartilage matrix remodeling and vascular invasion are required for the migration and differentiation of osteoclasts and osteoblasts, which remove the mineralized cartilage matrix and replace it with bone. DEVELOPMENT OF THE JOINT CAPSULE AND SYNOVIUM The interzone and the contiguous perichondrial envelope, of which the interzone is a part, contain the mesenchymal cell precursors that give rise to other joint components, including the joint capsule, synovial lining, menisci, intracapsular ligaments, and tendons.3,4,103,104 The external mesenchymal tissue condenses as a fibrous capsule. The peripheral mesenchyme becomes vascularized and is incorporated as the synovial mesenchyme, which differentiates into a pseudomembrane at about the same time as cavitation begins in the central interzone (stage 23, approximately 8 weeks). The menisci arise from the eccentric portions of the articular interzone. In common usage, the term synovium refers to the true synovial lining and the subjacent vascular and areolar tissue, up to—but excluding—the capsule. Synovial lining cells can be distinguished as soon as the multiple cavities within the interzone begin to coalesce. At first, these are exclusively fibroblast-like (type B) cells. As the joint cavity increases in size, synovial-lining cell layers expand by proliferation of fibroblast-like cells and recruitment of macrophage-like (type A) cells from the circulation.105 The synovial lining cells express the hyaluronan receptor, CD44, and UDPGD, the levels of which remain elevated after cavitation. This increased activity likely contributes to the high concentration of hyaluronan in joint fluids.31,106 Further synovial expansion results in the appearance of synovial villi at the end of the second month, early in the fetal period, which greatly increases the surface area available for exchange between the joint cavity and the vascular space. The role of innervation in the developing joint is not well understood. A dense capillary network develops in the subsynovial tissue, with numerous capillary loops that penetrate into the true synovial lining layer. The human synovial microvasculature is already innervated by 8 weeks (stage 23) of gestation, around the time of joint cavitation,103 as is shown by immunoreactivity for the neuronal “housekeeping” enzymes.107 Evidence of neurotransmitter function is not found until much later, however, with the appearance of the sensory neuropeptide, substance P, at 11 weeks. The putative sympathetic neurotransmitter, neuropeptide Y, appears at 13 weeks of gestation, along with the catecholamine­synthesizing enzyme tyrosine hydroxylase. The finding that the Slit2 gene, which functions for the guidance of neuronal axons and neurons, is expressed in the mesenchyme adjacent to the AER (stages 20 to 22) and in peripheral mesenchyme of the limb bud (stages 23 to 28) suggests that innervation is an integral part of synovial joint development.108 DEVELOPMENT OF NONARTICULAR JOINTS In contrast to articular joints, the temporomandibular joint develops slowly, with cavitation at a crown-rump length of 57 to 75 mm (i.e., well into the fetal stage).109 This slow







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development may be because this joint develops in the absence of a continuous blastema and involves the insertion between bone ends of a fibrocartilaginous disk that arises from muscular and mesenchymal derivatives of the first pharyngeal arch. The development of other types of joints, such as synarthroses, is similar to that of diarthrodial joints except that cavitation does not occur, and synovial mesenchyme is not formed. In these respects, synarthroses and amphiarthroses resemble the “fused” peripheral joints induced by paralyzing chicken embryos,110 and they may develop as they do because there is relatively little motion during their formation. Human vertebrae and intervertebral disks develop as units, each derived from a homogeneous blastema arising from a somite. Each embryonic intervertebral disk serves as a rostral and caudal chondrogenic zone for the two adjacent evolving vertebral bodies. The periphery of the embryonic “disk” is replaced by the anulus fibrosus.111 The intervertebral disk bears many similarities to the joint; the anulus is the joint capsule, the nucleus pulposus is the joint cavity, and the vertebral end plates are the cartilage-covered bone ends composing the articulation. The proteoglycans and collagens expressed during development of the intervertebral disk have been mapped and reflect the complex structure-function relationships that allow flexibility and resistance to compression in the spine.112-115 DEVELOPMENT OF ARTICULAR CARTILAGE In the vertebrate skeleton, cartilage is the product of cells from three distinct embryonic lineages. Craniofacial cartilage is formed from cranial neural crest cells, the cartilage of the axial skeleton (intervertebral disks, ribs, and sternum) forms from paraxial mesoderm (somites), and the articular cartilage of the limbs is derived from the lateral plate mesoderm.2 In the developing limb bud, mesenchymal condensations, followed by chondrocyte differentiation and maturation, occur in digital zones, whereas undifferentiated mesenchymal cells in the interdigital web zones undergo cell death.116 Embryonic cartilage is destined for one of several fates: It can remain as permanent cartilage, as on the articular surfaces of bones, or it can provide a template for the formation of bones by endochondral ossification. During development, chondrocyte maturation expands from the central site of the original condensation, which forms the cartilage anlage resembling the shape of the future bone, toward the ends of the forming bones. During joint cavitation, the peripheral interzone is absorbed into each adjacent cartilaginous zone, evolving into the articular surface. The articular surface is destined to become a specialized cartilaginous structure that does not normally undergo vascularization and ossification. More recent evidence indicates that postnatal maturation of the articular cartilage involves an appositional growth mechanism originating from progenitor cells at the articular surface, rather than by an interstitial mechanism.113 The chondrocytes of mature articular cartilage are terminally differentiated cells that continue to express cartilage-specific matrix molecules, such as type II collagen and aggrecan (see following section).19,21,24 Through the processes described previously, the articular joint spaces are developed and lined



on all surfaces either by cartilage or by synovial lining cells. These two different tissues merge at the enthesis, the region at the periphery of the joint where the cartilage melds into bone, and where ligaments and the capsule are attached.117 In the postnatal growth plate, the differentiation of the perichondrium also is linked to the differentiation of the chondrocytes in the epiphysis into different zones of the growth plate, and contributes to longitudinal bone growth.28,78



ORGANIZATION AND PHYSIOLOGY OF THE MATURE JOINT The unique structural properties and biochemical components of diarthrodial joints make them extraordinarily durable load-bearing devices.118 The mature diarthrodial joint is a complex structure, influenced by its environment and mechanical demands (see Chapter 6). There are structural differences between joints determined by their different functions. The shoulder joint, which demands an enormous range of motion, is stabilized primarily by muscles, whereas the hip, requiring motion and antigravity stability, has an intrinsically stable ball-and-socket configuration. The components of the “typical” synovial joint are the synovium, muscles, tendons, ligaments, bursae, menisci, articular cartilage, and subchondral bone. The anatomy and physiology of muscles are described in detail in Chapter 5. SYNOVIUM The synovium lines the joint cavity and is the sight of production of synovial fluid that provides the nutrition for the articular cartilage and lubricates the cartilage surfaces. It is a thin membrane between the fibrous joint capsule and fluid-filled synovial cavity that attaches to skeletal tissues at the bone-cartilage interface and does not encroach on the surface of the articular cartilage. It is divided into functional compartments: the lining region (synovial intima), the subintimal stroma, and the neurovasculature (Fig. 1-6). The synovial intima, also termed synovial lining, is the superficial layer of the normal synovium that is in contact with the intra-articular cavity.106,119 The synovial lining is loosely attached to the subintima, which contains blood vessels, lymphatics, and nerves. Capillaries and arterioles generally are located directly underneath the synovial intima, whereas venules are located closer to the joint capsule. A transition from loose to dense connective tissue occurs from the joint cavity to the capsule. Most cells in the normal subintimal stroma are fibroblasts and macrophages, although adipocytes and occasional mast cells are present.106 These compartments are not circumscribed by basement membranes, but nonetheless have distinct functions; they are separated from each other by chemical barriers, such as membrane peptidases, which limit the diffusion of regulatory factors between compartments. Synovial compartments are unevenly distributed within a single joint. Vascularity is high at the enthesis where synovium, ligament, and cartilage coalesce.120 Far from being a homogeneous tissue in continuity with the synovial cavity, synovium is highly heterogeneous, and synovial fluid may be poorly representative of the tissue-fluid composition of any synovial tissue compartment. In rheumatoid arthritis, the synovial lining of diarthrodial joints is the site of the initial inflammatory



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Intimal macrophage



Intimal fibroblasts



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STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE







Synovial fluid



Subintimal fibroblast Subintimal macrophage Blood vessels



A



B



Figure 1-6  A, Schematic representation of normal human synovium. The intima contains specialized fibroblasts expressing vascular cell adhesion molecule-1 (VCAM-1) and uridine diphosphoglucose (UDPG) and specialized macrophages expressing FcγRIIIa. The deeper subintima contains ­unspecialized counterparts. B, Microvascular endothelium in human synovium contains receptors for the vasodilator/growth factor substance P. Silver grains represent specific binding of [125I]Bolton Hunter–labeled substance P to synovial microvessels (arrows). Arrowheads indicate the synovial surface. Emulsion-dipped in vitro receptor autoradiography preparations with hematoxylin and eosin counterstain. Calibration bar = 1 μm. (A from Edwards JCW: Fibroblast biology: Development and differentiation of synovial fibroblasts in arthritis. Arthritis Res 2:344-347, 2000.)



process.121,122 This lesion is characterized by proliferation of the synovial lining cells, increased vascularization, and infiltration of the tissue by inflammatory cells, including lymphocytes, plasma cells, and activated macrophages (see Chapter 65).123-125 Synovial Lining The synovial lining, a specialized condensation of mesenchymal cells and extracellular matrix, is located between the synovial cavity and stroma. In normal synovium, the lining layer is two to three cells deep, although intra-articular fat pads usually are covered by only a single layer of synovial cells, and ligaments and tendons are covered by synovial cells that are widely separated. At some sites, lining cells are absent, and the extracellular connective tissue constitutes the lining layer.126 Such “bare areas” become increasingly frequent with advancing age.127 Although the synovial lining is often referred to as the synovial membrane, the term membrane is more correctly reserved for epithelia that have basement membranes, tight intercellular junctions, and desmosomes. Instead, synovial lining cells lie loosely in a bed of hyaluronate interspersed with collagen fibrils. This is the macromolecular sieve that imparts the semipermeable nature of the synovium. The absence of any true epithelial tissue, including basement membrane, is a major determinant of joint physiology. Early electron microscopic studies characterized lining cells as macrophage-derived type A synoviocytes and fibroblast-derived type B synoviocytes.128 High UDPGD activity and CD55 are used to distinguish type B synovial cells, whereas nonspecific esterase and CD68 typify type A cells.129,130 Normal synovium is lined predominantly by fibroblast-like cells, whereas macrophage-like cells compose only 10% to 20% of lining cells (see Fig. 1-6). Type A, macrophage-like synovial cells contain vacuoles, a prominent Golgi apparatus, and filopodia, but they have little rough endoplasmic reticulum. These cells express numerous cell surface markers of the monocyte-macrophage lineage, including CD11b, CD68, CD14, CD163, and the



IgG Fc receptor, FcγRIIIa.106 Synovial intimal macrophages are phagocytic and may provide a mechanism by which particulate matter can be cleared from the normal joint cavity. Similar to other tissue macrophages, these cells have little capacity to proliferate and are likely localized to the joint during development. The op/op osteopetrotic mouse that is deficient in macrophages because of an absence of macrophage colony-stimulating factor also lacks synovial macrophages.131 This finding provides further evidence that type A synovial cells are of a common lineage with other tissue macrophages. Although they represent only a small percentage of the cells in the normal synovium, the macrophages are recruited from the circulation during synovial inflammation, partly from subchondral bone marrow through vascular channels near the enthesis. The type B, fibroblast-like synovial cell contains fewer vacuoles and filopodia than type A cells and has abundant protein-synthetic organelles. Similar to other fibroblasts, lining cells express the collagen synthesis enzyme prolyl hydroxylase and synthesize extracellular matrix components, including collagens, sulfated proteoglycans, fibronectin, fibrillin-1, and tenascin.106,132 They have the potential to proliferate, although proliferation markers are rarely seen in normal synovium.133 In contrast to stromal fibroblasts, synovial intimal fibroblasts express UDPGD and synthesize hyaluronan, an important constituent of synovial fluid.106 They also synthesize lubricin, which, together with hyaluronan, is necessary for the low-friction interaction of cartilage surfaces in the diarthrodial joint. Despite not being a true epithelium, synovial lining cells bear abundant membrane peptidases on their surface, capable of degrading a wide range of regulatory peptides, such as substance P and angiotensin II.134 These enzymes may be important in limiting the diffusion of these potent peptide mediators away from the immediate vicinity of their site of release and action. Normal synovial lining cells also express a rich array of adhesion molecules, including CD44, the principal receptor for hyaluronan; vascular cell adhesion molecule (VCAM)-1; and intercellular adhesion molecule (ICAM)-1.106,135-137 These are probably essential for cellular attachment to



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specific matrix components in the synovial lining region, preventing loss into the synovial cavity of cells subjected to deformation and shear stresses during joint movement. Adhesion molecules such as VCAM-1 and ICAM-1 potentially also are involved in the recruitment of inflammatory cells during the evolution of arthritis. Cadherins mediate cell-cell adhesion between adjacent cells of the same type. The identification of cadherin-11 as a key adhesion molecule that regulates the formation of the synovial lining during development and the synoviocyte function postnatally has provided the opportunity to examine its role in inflammatory joint disease.138 Cadherin-11 deficiency or treatment with cadherin-11 antibody or a cadherin-11 fusion protein reduced synovial inflammation and reduced cartilage erosion in an animal model of arthritis.139 Synovial Vasculature The subintimal synovium contains blood vessels, providing the blood flow that is required for solute and gas exchange in the synovium itself and for the generation of synovial fluid.120 The avascular articular cartilage also depends on nutrition in the synovial fluid, derived from the synovial vasculature. The vascularized synovium behaves similar to an endocrine organ, generating factors that regulate synoviocyte function and serving as a selective gateway that recruits cells from the circulation during stress and inflammation.140 Finally, synovial blood flow plays an important role in regulating intra-articular temperature. The synovial vasculature can be divided, on morphologic and functional grounds, into arterioles, capillaries, and venules. In addition, lymphatics accompany arterioles and larger venules.106,120 Arterial and venous networks of the joint are complex and are characterized by arteriovenous anastomoses that communicate freely with blood vessels in periosteum and periarticular bone. As large synovial arteries enter the deep layers of the synovium near the capsule, they give off branches, which bifurcate again to form “microvascular units” in the subsynovial layers. The synovial lining region, the surfaces of intra-articular ligaments, and the entheses (in the angle of ligamentous insertions into bone) are particularly well vascularized.120 The distribution of synovial vessels, which were formed largely as a result of vasculogenesis during development of the joint, displays considerable plasticity. Vasculogenesis is a dynamic process that depends on the cellular interactions with regulatory factors and the extracellular matrix that are also important in angiogenesis. In inflammatory arthritis, the density of blood vessels decreases relative to the growing synovial mass, creating a hypoxic and acidotic environment.141,142 Angiogenic factors such as VEGF, acting via VEGF receptor 1 and 2 (Flt-1 and Flk-1), and basic FGF promote proliferation and migration of endothelial cells, a process that is facilitated by matrix-degrading enzymes and adhesion molecules such as integrin αvβ3 and E-selectin, expressed by activated endothelial cells.143-145 Vessel maturation is facilitated by angiopoietin-1 acting via the Tie-2 receptor. The angiogenic molecules are restricted to the capillary epithelium in normal synovium, but their levels are elevated in inflamed synovium in perivascular sites and areas remote from vessels.146,147



Regulation of Synovial Blood Flow Synovial blood flow is regulated by intrinsic (autocrine and paracrine) and extrinsic (neural and humoral) systems. Locally generated factors, such as the peptide vasoconstrictors angiotensin II and endothelin-1, act on adjacent arteriolar smooth muscle to regulate regional vascular tone.120 Normal synovial arterioles are richly innervated by sympathetic nerves containing vasoconstrictors, such as norepinephrine and neuropeptide Y, and by “sensory” nerves that also play an efferent vasodilatory role by releasing neuropeptides, such as substance P and calcitonin gene–related peptide.148,149 Arterioles regulate regional blood flow. Capillaries and postcapillary venules are sites of fluid and cellular exchange. Correspondingly, regulatory systems are differentially distributed along the vascular axis. Angiotensin-converting enzyme, which generates angiotensin II, is localized predominantly in arteriolar and capillary endothelia and decreases during inflammation.150 Specific receptors for angiotensin II and for substance P are abundant on synovial capillaries, with lower densities on adjacent arterioles. Dipeptidyl peptidase IV, a peptide-degrading enzyme, is specifically localized to the cell membranes of venular endothelium. The synovial vasculature is not only functionally compartmentalized from the surrounding stroma, but also highly specialized along its arteriovenous axis. Other unique characteristics of the normal synovial vasculature include the presence of inducible nitric oxidase synthase–independent 3-nitrotyrosine, a reaction product of peroxynitrite,151 and the localization of the synoviocyte-derived CXCL12 chemokine on heparan sulfate receptors on endothelial cells,152 suggesting physiologic roles for these molecules in normal vascular function. JOINT INNERVATION Dissection studies have shown that each joint has a dual nerve supply, consisting of specific articular nerves that penetrate the capsule as independent branches of adjacent peripheral nerves and articular branches that arise from related muscle nerves. The definition of joint position and the detection of joint motion are monitored separately and by a combination of multiple inputs from different receptors in varied systems. Nerve endings in muscle and skin and in the joint capsule mediate sensation of joint position and movement.153,154 Normal joints have afferent (sensory) and efferent (motor) innervations. Fast-conducting, myelinated A fibers innervating the joint capsule are important for proprioception and detection of joint movement; slow­conducting, unmyelinated C fibers transmit diffuse pain sensation and regulate synovial microvascular function. Normal synovium is richly innervated by fine, unmyelinated nerve fibers that follow the courses of blood vessels and extend into the synovial lining layers.148 These nerve fibers do not have specialized endings and are slow-conducting fibers; they may transmit diffuse, burning, or aching pain sensation. Sympathetic nerve fibers surround blood vessels, particularly in the deeper regions of normal synovium. They contain and release classic neurotransmitters, such as norepinephrine, and neuropeptides that constrict synovial blood vessels. Neuropeptides that are markers of sensory nerves include substance P, calcitonin gene–related peptide, neuropeptide Y, and vasoactive intestinal peptide.148,155-157



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Afferent nerves containing substance P also have an efferent role in the synovium. Substance P is released from peripheral nerve terminals into the joint, and specific, G protein–coupled receptors for substance P are localized to microvascular endothelium in normal synovium. Abnormalities of articular innervation that are associated with inflammatory arthritis may contribute to the failure of synovial inflammation to resolve.148,158 Excessive local neuropeptide release may result in the loss of nerve fibers owing to neuropeptide depletion. Synovial tissue proliferation without concomitant growth of new nerve fibers may lead to an apparent partial denervation of synovium.148,158 Studies in patients suggest that free nerve endings containing substance P may modulate inflammation and the pain pathway in osteoarthritis.159 Afferent nerve fibers from the joint play an important role in the reflex inhibition of muscle contraction. Trophic factors generated by motoneurons, such as the neuropeptide calcitonin gene–related peptide, are important in maintaining muscle bulk and a functional neuromuscular junction.160 Decreases in motoneuron trophic support during articular inflammation probably contribute to muscle wasting. Mechanisms of joint pain have been reviewed in detail.161,162 In a noninflamed joint, most sensory nerve fibers do not respond to movement within the normal range; these are referred to as silent nociceptors. In an acutely inflamed joint, however, these nerve fibers become sensitized by mediators, such as bradykinin, neurokinin 1, and prostaglandins (peripheral sensitization), such that normal movements induce pain. Pain sensation is upregulated or downregulated further in the central nervous system, at the level of the spinal cord and in the brain, by central sensitization and “gating” of nociceptive input. Although the normal joint may respond predictably to painful stimuli, there is often a poor correlation between apparent joint disease and perceived pain in chronic arthritis. Pain associated with joint movements within the normal range is a characteristic symptom described by patients with chronically inflamed joints caused by rheumatoid arthritis. Chronically inflamed joints may not be painful at rest, however, unless acutely inflamed.163 TENDONS Tendons are functional and anatomic bridges between muscle and bone.164,165 They focus the force of a large mass of muscle into a localized area on bone and, by splitting to form numerous insertions, may distribute the force of a single muscle to different bones. Tendons are formed of longitudinally arranged collagen fibrils embedded in an organized, hydrated proteoglycan matrix with blood vessels, lymphatics, and fibroblasts.166 Cross-links between adjacent collagen chains or molecules contribute to the tensile strength of the tendon. 167,168 Tendon collagen fibrillogenesis is initiated during early development by a highly ordered process of alignment involving the actin cytoskeleton and cadherin-11.169,170 Many tendons, particularly tendons with a large range of motion, run through vascularized, discontinuous sheaths of collagen lined with mesenchymal cells resembling synovium. Gliding of tendons through their sheaths is enhanced by hyaluronic acid produced by the lining cells. Tendon movement is essential for the embryogenesis and



11



maintenance of tendons and their sheaths. Degenerative changes appear in tendons, and fibrous adhesions form between tendons and sheaths when inflammation or surgical incision is followed by long periods of immobilization.171 At the myotendinous junction, recesses between muscle cell processes are filled with collagen fibrils, which blend into the tendon. At its other end, collagen fibers of the tendon typically blend into fibrocartilage, mineralize, and merge into bone through a fibrocartilaginous transition zone termed the enthesis, or insertion site.172 Tendon fibroblasts synthesize and secrete collagens, proteoglycans, and other matrix components, such as fibronectin and tenascin C, and MMPs and their inhibitors, which can contribute to the breakdown and repair of tendon components.166,173-176 Collagen fibrils in tendon are composed primarily of type I collagen with some type III collagen, but there are regional differences in the distribution of other matrix components. The compressed region contains the small proteoglycans, biglycan, decorin, fibromodulin, and lumican, and the large proteoglycan versican.177,178 The major components in the tensile region of the tendon are decorin, microfibrillar type VI collagen, fibromodulin, and proline and arginine-rich end leucine-rich repeat protein. The presence of cartilage oligomeric matrix protein, aggrecan, and biglycan and collagen types II, IX, and XI is indicative of fibrocartilage.179,180 The collagen fiber orientation at the tendon-to-bone enthesis is important for maintaining microarchitecture, by reducing the stress concentrations and shielding the outward splay of the insertion from the highest stresses.181 Understanding the structure has implications for tendon repair because motion between a tendon graft and bone tunnel may impair early graft incorporation and lead to tunnel widening secondary to bone resorption.182 Failure of the muscle-tendon apparatus is rare, but when it does occur, it is secondary to enormous, quickly generated forces across a joint and usually occurs near the tendon insertion into bone.183,184 Factors that may predispose to tendon failure are aging processes, including loss of extracellular water and an increase in intermolecular cross-links of collagen; tendon ischemia; iatrogenic factors, including injection of glucocorticoids; and deposition of calcium hydroxyapatite crystals within the collagen bundles. Alterations in collagen fibril composition and structure are associated with tendon degeneration during aging and may predispose to osteoarthritis.185,186 Evidence indicates that BMPs promote tendon repair if osteogenic signaling is impaired.187 LIGAMENTS Ligaments provide a stabilizing bridge between bones, permitting a limited range of movement.188 The ligaments often are recognized only as hypertrophied components of the fibrous joint capsule and are structurally similar to tendons.189 Although the fibers are oriented parallel to the longitudinal axis of both tissues,164 the collagen fibrils in ligaments are nonparallel and arranged in fibers that are oriented roughly along the long axis in a wavy, undulating pattern, or “crimp,” which can straighten in response to load. Some ligaments have a higher ratio of elastin to collagen (1:4) than tendons (1:50), which permits a greater degree of stretch. Ligaments also have larger amounts of reducible cross-links, more type III collagen, slightly less total ­collagen,



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and more glycosaminoglycans compared with tendons. The cells in ligaments seem to be more metabolically active than the cells in tendons because they have more plump cellular nuclei and higher DNA content. During postnatal growth, the development of ligament attachment zones involves changes in the ratios and distribution of collagen types I, III, and V and the synthesis of type II collagen and proteoglycans by fibrochondrocytes that develop from ligament cells at the attachment zone.190,191 Attachment zones are believed to permit gradual transmission of the tensile force between ligament and bone. Ligaments play a major role in the passive stabilization of joints, aided by the capsule and, when present, menisci. In the knee, the collateral and cruciate ligaments provide stability when there is little or no load on the joint. As compressive load increases, there is an increasing contribution to stability from the joint surfaces themselves and the surrounding musculature. Injured ligaments generally heal, and structural integrity is restored by contracture of the healing ligament so that it can act again as a stabilizer of the joint.192 BURSAE The many bursae in the human body facilitate gliding of one tissue over another, much as a tendon sheath facilitates movement of its tendon. Bursae are closed sacs, lined sparsely with mesenchymal cells similar to synovial cells, but they are generally less well vascularized than synovium. Most bursae differentiate concurrently with synovial joints during embryogenesis. During life, however, trauma or inflammation may lead to the development of new bursae, hypertrophy of previously existing ones, or communication between deep bursae and joints. In patients with rheumatoid arthritis, communications may exist between the subacromial bursae and the glenohumeral joint, between the gastrocnemius or semimembranosus bursae and the knee joint, and between the iliopsoas bursa and the hip joint. It is unusual, however, for subcutaneous bursae, such as the prepatellar bursa or olecranon bursa, to develop communication with the underlying joint.193 MENISCI The meniscus, a fibrocartilaginous, wedge-shaped structure, is best developed in the knee, but also is found in the acromioclavicular and sternoclavicular joints, the ulnocarpal joint, and the temporomandibular joint.194,195 Until more recently, menisci were thought to have little function and a quiescent metabolism with no capability of repair, although early observations indicated that removal of menisci from the knee may lead to premature arthritic changes in the joint.196 Evidence from an arthroscopic study of patients with anterior cruciate ligament insufficiency indicates that the pathology of the medial meniscus correlates with that of the medial femoral cartilage.197 The meniscus is now considered to be an integral component of the knee joint that has important functions in joint stability, load distribution, shock absorption, and lubrication.194,195 The microanatomy of the meniscus is complex and age dependent.198 The characteristic shape of the lateral and the medial menisci is achieved early in prenatal



d­ evelopment. At that time, the menisci are cellular and highly vascularized; with maturation, vascularity decreases progressively from the central margin to the peripheral margin. After skeletal maturity, the peripheral 10% to 30% of the meniscus remains highly vascularized by a circumferential capillary plexus and is well innervated.199 Tears in this vascularized peripheral zone may undergo repair and remodeling.200 The central portion of the mature meniscus is an avascular fibrocartilage, however, without nerves or lymphatics, consisting of cells surrounded by an abundant extracellular matrix of collagens, chondroitin sulfates, dermatan sulfates, and hyaluronic acid. Tears in this central zone heal poorly, if at all. Collagen constitutes 60% to 70% of the dry weight of the meniscus and is mostly type I collagen, with lesser amounts of types III, V, and VI. A small quantity of cartilage-specific type II collagen is localized to the inner, avascular portion of the meniscus. Collagen fibers in the periphery are mostly circumferentially oriented, with radial fibers extending toward the central portion.201-204 Elastin content is around 0.6%, and proteoglycan content is around 2% to 3% dry weight. Aggrecan and decorin are the major proteoglycans in the adult meniscus.205,206 Decorin is the predominant proteoglycan synthesized in the meniscus from young individuals, whereas the relative proportion of aggrecan synthesis increases with age. Although the capacity of the meniscus to synthesize sulfated proteoglycans decreases after the teenage years, the age-related increases in expression of decorin and aggrecan mRNA suggest that the resident cells are able to respond quickly to alterations in the biomechanical environment.207 The meniscus was defined originally as a fibrocartilage, based on the rounded or oval shape of most of the cells and the fibrous microscopic appearance of the extracellular matrix.208 Based on molecular and spatial criteria, three distinct populations of cells are recognized in the meniscus of the knee joint202: 1. The fibrochondrocyte is the most abundant cell in the middle and inner meniscus, synthesizing primarily type I collagen and relatively small amounts of type II and III collagens. It is round or oval in shape and has a pericellular filamentous matrix containing type VI collagen. 2. The fibroblast-like cells lack a pericellular matrix and are located in the outer portion of the meniscus. They are distinguished by long, thin, branching cytoplasmic projections that stain for vimentin. They make contact with other cells in different regions via connexin 43–containing gap junctions. The presence of two centrosomes, one associated with a primary celium, suggests a sensory, rather than motile, function that could enable the cells to respond to circumferential tensile loads, rather than compressive loads.209 3. The superficial zone cells have a characteristic fusiform shape with no cytoplasmic projections. The occasional staining of these cells in the uninjured meniscus with α-actin and their migration into surrounding wound sites suggest that they are specialized progenitor cells that may participate in a remod­ eling response in the meniscus and surrounding tissues.210,211



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STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



MATURE ARTICULAR CARTILAGE Articular cartilage is a specialized connective tissue that covers the weight-bearing surfaces of diarthrodial joints.118,212,213 The principal functions of cartilage layers covering bone ends are to permit low-friction, high-velocity movement between bones, to absorb the transmitted forces associated with locomotion, and to contribute to joint stability. Lubrication by synovial fluid provides frictionless movement of the articulating cartilage surfaces. Chondrocytes (see Chapter 3) are the single cellular component of adult hyaline articular cartilage and are responsible for synthesizing and maintaining the highly specialized cartilage matrix macromolecules. The cartilage extracellular matrix is composed of an extensive network of collagen fibrils, which confers tensile strength, and an interlocking mesh of proteoglycans, which provides compressive stiffness through the ability to absorb and extrude water. Numerous other noncollagenous proteins also contribute to the unique properties of cartilage (Table 1-1). Histologically, the tissue appears to be fairly homogeneous and clearly distinguished from the calcified cartilage and underlying subchondral bone (Fig. 1-7). The organization of articular cartilage and structure-function relationships of cartilage matrix components are described in Chapter 3. SUBCHONDRAL BONE INTERACTIONS WITH ARTICULAR CARTILAGE The subchondral bone plate beneath the calcified base of articular cartilage may have many effects on the cartilage above it. Its stiffness modifies the compressive forces to which articular cartilage is subjected, its blood supply may be important in cartilage nutrition (see following section), and its cells may produce peptides that regulate chondrocyte function. Several studies have suggested that the responses of the subchondral bone to mechanical stimulation may transmit signals into the articular cartilage.214 Tidemark advancement with thickening of the calcified cartilage and thinning of articular cartilage is associated with fibrillation of the cartilage surface during aging.215 An increase in subchondral bone density is an early feature of osteoarthritis.216 Radin and Rose217 proposed that the initiation of fibrillation is caused by an increase in subchondral bone stiffness. These changes in subchondral bone stiffness may be secondary to cartilage deterioration, but are necessary for progression of osteoarthritic lesions, and involve bone and calcified cartilage close to the joint.218-220 Also, at the junction of the articular hyaline cartilage and adjacent subchondral bone, there is evidence of vascular invasion and advancement in the zone of calcified cartilage in the region of the so-called tidemark that contributes further to a decrease in articular cartilage thickness.221 A more recent study showed that angiogenesis in the osteochondral junction is independent of synovial angiogenesis and synovitis, but is associated with cartilage changes and clinical disease activity.222 These structural alterations in the articular cartilage and periarticular bone may lead to modification of the contours of the adjacent articulating surfaces.217,223-225 Because increased trabecular bone volume with trabecular sclerosis and increased bone turnover are features of osteoarthritis pathogenesis, therapies that target bone have been proposed. Examples are calcitonin,226,227



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b­ isphosphonates,228 and estrogen.229 Although receptor activator of nuclear factor ĸB (NFĸB) ligand (RANKL), which mediates osteoclast differentiation and activity, and its receptor RANK, a member of the tumor necrosis factor receptor family, are expressed in adult articular chondrocytes, exogenous RANKL does not activate NFĸB or stimulate the production of collagenase or nitric oxide.230 Inhibition of RANKL expression does not block cartilage destruction in ­ inflammatory models,231 although RANKL may have indirect effects on cartilage by its protective effect on bone.232 Table 1-1  Extracellular Matrix Components   of Articular Cartilage* Collagens Type II Type IX Type XI Type VI Types XII, XIV Type X (hypertrophic chondrocyte) Proteoglycans Aggrecan Versican Link protein Biglycan (DS-PGI) Decorin (DS-PGII) Epiphycan (DS-PGIII) Fibromodulin Lumican Proline/arginine-rich and leucine-rich repeat protein (PRELP) Chondroadherin Perlecan Lubricin (SZP) Other Noncollagenous Proteins (Structural) Cartilage oligomeric matrix protein (COMP) or thrombospondin-5 Thrombospondin-1 and thrombospondin-3 Cartilage matrix protein (matrilin-1) and matrilin-3 Fibronectin Tenascin-C Cartilage intermediate layer protein (CILP) Fibrillin Elastin Other Noncollagenous Proteins (Regulatory) Glycoprotein (gp)-39, YKL-40 Matrix Gla protein (MGP) Chondromodulin-I (SCGP) and chondromodulin-II Cartilage-derived retinoic acid–sensitive protein (CD-RAP) Growth factors Cell Membrane–Associated Proteins Integrins (α1β1, α2β1, α3β1, α5β1, α6β1, α10β1, αvβ3, αvβ5) Anchorin CII (annexin V) Cell determinant 44 (CD44) Syndecan-3 Discoidin domain receptor 2 *The collagens, proteoglycans, and other noncollagenous proteins in the cartilage matrix are synthesized by chondrocytes at different stages during development and growth of cartilage. In mature articular cartilage, proteoglycans and other noncollagen proteins are turned over slowly, whereas the collagen network is stable unless exposed to proteolytic cleavage. Proteins that are associated with chondrocyte cell membranes also are listed because they permit specific interactions with extracellular matrix proteins. The specific structure-function relationships are discussed in Chapter 3 and described in Table 3-1. DS-PG, dermatan sulfate proteoglycan; SCGP, small cartilage–derived glycoprotein; SZP, superficial zone protein; YKL-40, 40KD chitinase 3-like glycoprotein.



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Figure 1-7  A and B, Representative sections of normal human adult articular cartilage, showing nearly the same field in plain (A) and polarized (B) light. Note the clear demarcation of the articular cartilage from the calcified cartilage below the tidemark and the underlying subchondral bone.  (Hematoxylin-eosin stain; original magnification ×60.) (Courtesy of Edward F. DiCarlo, MD, Pathology Department, Hospital for Special Surgery, New York, NY.)



A



SYNOVIAL FLUID AND NUTRITION OF JOINT STRUCTURES The volume and composition of synovial fluid are determined by the properties of the synovium and its vasculature. Fluid in normal joints is present in small quantities (2.5 mL in the normal knee) sufficient to coat the synovial surface, but not to separate one surface from the other. Tendon sheath fluid and synovial fluid are biochemically similar. Both are essential for the nutrition and lubrication of adjacent avascular structures, including tendon and articular cartilage, and for limiting adhesion formation, maintaining movement. Characterization and measurement of synovial fluid constituents have proved useful for the identification of locally generated regulatory factors, markers of cartilage turnover, and the metabolic status of the joint, and for the assessment of the effects of therapy on cartilage homeostasis. Interpretation of such data requires, however, an understanding of the generation and clearance of synovial fluid and its various components. GENERATION AND CLEARANCE OF SYNOVIAL FLUID Synovial fluid concentrations of a protein represent the net contributions of synovial blood flow, plasma concentration, microvascular permeability, and lymphatic removal and its production and consumption within the joint space. Synovial fluid is a mixture of a protein-rich ultrafiltrate of plasma and hyaluronan synthesized by synoviocytes. Generation of this ultrafiltrate depends on the difference between intracapillary and intra-articular hydrostatic pressures and between colloid osmotic pressures of capillary plasma and synovial tissue fluid. Fenestrations, small pores covered by a thin membrane, in the synovial capillaries and the macromolecular sieve of hyaluronic acid facilitate rapid exchange



B



of small molecules, such as glucose and lactate, assisted—in the case of glucose—by an active transport system.233 Proteins are present in synovial fluid at concentrations inversely proportional to molecular size, with synovial fluid albumin concentrations being about 45% of those in plasma (Fig. 1-8).234 Concentrations of electrolytes and small molecules are equivalent to those in plasma.235 Synovial fluid is cleared through lymphatics in the synovium, assisted by joint movement. In contrast to ultrafiltration, lymphatic clearance of solutes is independent of molecular size. In addition, constituents of synovial fluid, such as regulatory peptides, may be degraded locally by enzymes, and low-molecular-weight metabolites may diffuse along concentration gradients into plasma. The kinetics of delivery and removal of a protein must be determined (e.g., using albumin as a reference solute) to assess the significance of its concentration in the joint.236 Hyaluronic acid is synthesized by fibroblast-like synovial lining cells, and it appears in high concentrations in synovial fluid at around 3 g/L, compared with a plasma concentration of 30 μg/L. Lubricin, a glycoprotein that assists articular lubrication, is another constituent of synovial fluid that is generated by the lining cells.363 It is now believed that hyaluronan functions in fluid-film lubrication, whereas lubricin is the true boundary lubricant in synovial fluid (see following). Because the volume of synovial fluid is determined by the amount of hyaluronan, water retention seems to be the major function of this large molecule.233,237 Despite the absence of a basement membrane, synovial fluid does not mix freely with extracellular synovial tissue fluid. Hyaluronan may trap molecules within the synovial cavity by acting as a filtration screen on the surface of the synovial lining, resisting the movement of synovial fluid out from the joint space.237 Synovial fluid and its constituent proteins have a rapid turnover time (around 1 hour



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STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



Probably RA



1.0



Gout Osteoarthritis Classic RA



Ratio



Normal



SF .10 S Conc.



Oroso- Trans- Cerulomucoid ferrin plasmin



44



74



160



α2 Macroglobulin



820



.01 1



100



1000



Molecular Weight (×103) Figure 1-8  Ratio of the concentration of proteins in synovial fluid to that found in serum, plotted as a function of molecular weight. Larger proteins are selectively excluded from normal synovial fluid, but this macromolecular sieve is less effective in diseased synovium. Conc., concentration; RA, rheumatoid arthritis; S, serum; SF, synovial fluid. (From Kushner I, Somerville JA: Permeability of human synovial membrane to ­plasma proteins. Arthritis Rheum 14:560, 1971. Reprinted with permission of the American College of Rheumatology.)



in normal knees), and equilibrium is not usually reached among all parts of the joint. Tissue fluid around fenestrated endothelium reflects plasma ultrafiltrate most closely, with a low content of hyaluronate compared with synovial fluid. Alternatively, locally generated or released peptides, such as endothelin and substance P, may attain much higher perivascular concentrations than those measured in synovial fluid. The turnover time for hyaluronan in the normal joint (13 hours) is an order of magnitude slower, however, than that of small solutes and proteins. Association with hyaluronan may result in trapping of solutes within synovial fluid.238 In normal joints, intra-articular pressures are slightly subatmospheric at rest (0 to −5 mm Hg).239 During exercise, hydrostatic pressure in the normal joint may decrease further. Resting intra-articular pressures in rheumatoid joints are around 20 mm Hg, whereas during isometric exercise, they may increase to greater than 100 mm Hg, well above capillary perfusion pressure and, at times, above arterial pressure. Repeated mechanical stresses can interrupt synovial perfusion during joint movement, particularly in the presence of a synovial effusion. SYNOVIAL FLUID AS AN INDICATOR OF JOINT FUNCTION In the absence of a basement membrane separating synovium or cartilage from synovial fluid, measurements made on synovial fluid may reflect the activity of these structures. A wide range of regulatory factors and products



15



of ­synoviocyte metabolism and cartilage breakdown may be generated locally within the joint, resulting in marked differences between the composition of synovial fluid and plasma ultrafiltrate. Because there is little capacity for the selective concentration of solutes in synovial fluid, solutes present at higher concentrations than in plasma are probably synthesized locally. It is necessary to know the local clearance rate, however, to determine whether the solutes present in synovial fluid at lower concentrations than in plasma are generated locally.235 Although microvascular permeability to protein in highly inflamed rheumatoid joints is more than twice that in osteoarthritic joints, synovial fluid protein concentrations vary little between the two joint diseases240 because the enhanced entry of proteins through the microvasculature is largely offset by the increased lymphatic clearance.241 Because clearance rates from synovial fluid may be slower than those from plasma, however, synovial fluid levels of drugs or urate may remain elevated after plasma levels have declined.233 Comparisons of synovial fluid constituents between disease groups are often limited by the sparseness of data on normal synovial fluid as a result of difficulties in its collection. Extrapolation from synovial fluid concentrations to local synthetic rates is complicated further because of variations in clearance rates and in synovial fluid volume. Plasma proteins are less effectively filtered in inflamed synovium, perhaps because of increased size of endothelial cell fenestrations or because interstitial hyaluronate-protein complexes are fragmented by enzymes associated with the inflammatory process.234 Concentrations of proteins, such as α2-macroglobulin (the principal proteinase inhibitor of plasma), fibrinogen, and IgM, are elevated in inflammatory synovial fluids (see Fig. 1-8), as are associated protein-bound cations. Membrane peptidases may limit the diffusion of regulatory peptides from their sites of release into synovial fluid. In inflammatory arthritis, fibrin deposits may retard flow between the tissue and the liquid phase. The cautious interpretation of synovial fluid analysis has important implications in understanding how to use data on biomarkers of cartilage damage and repair in rheumatoid arthritis and osteoarthritis (see Chapter 48). LUBRICATION AND NUTRITION OF THE ARTICULAR CARTILAGE Lubrication Synovial fluid serves as a lubricant for articular cartilage and a source of nutrition for the chondrocytes within. Lubrication is essential for protecting cartilage and other joint structures from friction and shear stresses associated with movement under loading. There are two basic categories of joint lubrication. In fluid-film lubrication, cartilage surfaces are separated by an incompressible fluid film; hyaluronan functions as the lubricant. In boundary lubrication, specialized molecules attached to the cartilage surface permit surface-to-surface contact, while decreasing the coefficient of friction. During loading, a noncompressible fluid film is trapped between opposing cartilage surfaces and prevents the surfaces from touching. Irregularities in the cartilage surface and its deformation during compression may augment this



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trapping of fluid. This stable film is approximately 0.1 μm thick in the normal human hip joint, but it can be much thinner in the presence of inflammatory synovial fluids or with increased cartilage porosity.242,243 Lubricin is the major boundary lubricant in the human joint.244 It is a glycoprotein, also called superficial zone protein and proteoglycan 4, which is synthesized by synovial cells and chondrocytes.245-248 It has a molecular weight of 225,000, is 200 nm in length, and is 1 to 2 nm in diameter.249 Dipalmitoyl phosphatidylcholine, which constitutes 45% of the lipid in normal synovial fluid, acts together with lubricin as a boundary lubricant.250 More recent work indicates that lubricin functions as a phospholipid carrier via a mechanism that is common to all tissues.251,252 Lipid composes 1% to 2% of dry weight of cartilage,253 and experimental treatment of cartilage surfaces with fat solvents impairs lubrication qualities.254 Nutrition As observed by Hunter in 1743,255 normal adult articular cartilage contains no blood vessels. Vascularization of cartilage would be expected to alter its mechanical properties. Blood flow would be repeatedly occluded during weight bearing and exercise, with reactive oxygen species generated during reperfusion, resulting in repeated damage to cartilage matrix and chondrocytes. Chondrocytes synthesize specific inhibitors of angiogenesis that maintain articular cartilage as an avascular tissue.256-258 As a result of the lack of adjacent blood vessels, the chondrocyte normally lives in an hypoxic and acidotic environment, with extracellular fluid pH values around 7.1 to 7.2,259 and it uses anaerobic glycolysis for energy production.260 High lactate levels in normal synovial fluid, compared with paired plasma measurements, partially reflect this anaerobic metabolism.261 There are two sources of nutrients for articular cartilage: (1) the synovial fluid and (2) subchondral blood vessels. The synovial fluid and, indirectly, the synovial lining, through which synovial fluid is generated, are the major sources of nutrients for articular cartilage. Nutrients may enter cartilage from synovial fluid either by diffusion or by mass transport of fluid during compression-relaxation cycles.262 Molecules as large as hemoglobin (65 kD) can diffuse through normal articular cartilage,263 and the solutes needed for cellular metabolism are much smaller. Diffusion of uncharged small solutes, such as glucose, is not impaired in matrices containing large amounts of glycosaminoglycans, and diffusivity of small molecules through hyaluronate is enhanced.264,265 Intermittent compression may serve as a pump mechanism for solute exchange in cartilage. The concept has arisen from observations that joint immobilization or dislocation leads to degenerative changes. In contrast, exercise increases solute penetration into cartilage in experimental systems.263 During weight bearing, fluid escapes from the load-bearing region by flow to other cartilage sites. When the load is removed, cartilage re-expands and draws back fluid, exchanging nutrients with waste materials.266 In a growing child, the deeper layers of cartilage are vascularized, such that blood vessels penetrate between columns of chondrocytes in the hypertrophic zone of the growth plate. It is likely that nutrients diffuse from these



tiny end capillaries through the matrix to chondrocytes. Diffusion from subchondral blood vessels is not considered a major route for the nutrition of normal adult articular cartilage because of the barrier provided by its densely calcified lower layer, the “tidemark.” Nonetheless, partial defects may normally exist in this barrier,267 and in arthritis, neovascularization of the deeper layers of articular cartilage may contribute to cartilage nutrition and to entry of inflammatory cells and cytokines.221,268 In aging and osteoarthritis, tidemark “duplication” may indicate communication between the bone and cartilage.218,269 Experimental studies have indicated that cartilage lesions of chondromalacia may develop if the subchondral blood supply of the patella is compromised.270



SUMMARY AND CONCLUSION Normal human synovial joints are complex structures that comprise interacting connective tissue elements that permit constrained and low-friction movement of adjacent bones. The development of synovial joints in the embryo is a highly ordered process involving complex cell-cell and cell-matrix interactions that lead to the formation of the cartilage anlage and interzone and joint cavitation. Understanding of the cellular interactions and molecular factors involved in cartilage morphogenesis and limb development has provided clues to understanding the functions of the synovium, articular cartilage, and associated structures in the mature joint. The synovial joint is uniquely adapted to responding to environmental and mechanical demands. The synovial lining is composed of two to three cell layers, and there is no basement membrane separating the lining cells from the underlying connective tissue. The synovium produces synovial fluid, which provides nutrition and lubrication to the avascular articular cartilage. Normal articular cartilage contains a single cell type, the articular chondrocyte, which is responsible for maintaining the integrity of the extracellular cartilage matrix. This matrix consists of a complex network of collagens, proteoglycans, and other noncollagenous proteins, which provide tensile strength and compressive resistance. Proper distribution and relative composition of these proteins is required for the function of cartilage in protecting the subchondral bone from adverse environmental influences. Maintenance of the unique composition and organization of each joint tissue is crucial for normal joint function, which is compromised in response to inflammation, biomechanical injury, and aging. Knowledge of the normal structure-function relationships within joint tissues is essential for understanding the pathogenesis and consequences of joint diseases.



REFERENCES   1. Simkin PA: The muskuloskeletal system, A: Joints. In Klippel JH, Crofford LJ, Stone JH, et al (eds): Primer on the Rheumatic Diseases. Atlanta, Arthritis Foundation, 2001, pp 5-9.   2. Olsen BR, Reginato AM, Wang W: Bone development. Annu Rev Cell Dev Biol 16:191-220, 2000.   3. O’Rahilly R, Gardner E: The timing and sequence of events in the development of the limbs in the human embryo. Anat Embryol (Berl) 148:1-23, 1975.



PART 1 



| 



STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



  4. O’Rahilly R, Gardner E: The embryology of movable joints. In Sokoloff L (ed): The Joints and Synovial Fluid, Vol 1. New York, Academic Press, 1978, p 49.   5. Pacifici M, Koyama E, Iwamoto M: Mechanisms of synovial joint and articular cartilage formation: Recent advances, but many lingering mysteries. Birth Defects Res C Embryo Today 75:237-248, 2005.   6. Church VL, Francis-West P: Wnt signalling during limb development. Int J Dev Biol 46:927-936, 2002.   7. Francis-West PH, Abdelfattah A, Chen P, et al: Mechanisms of GDF5 action during skeletal development. Development 126:1305-1315, 1999.   8. Edwards CJ, Francis-West PH: Bone morphogenetic proteins in the development and healing of synovial joints. Semin Arthritis Rheum 31:33-42, 2001.   9. Xu X, Weinstein M, Li C, et al: Fibroblast growth factor receptors (FGFRs) and their roles in limb development. Cell Tissue Res 296:33-43, 1999. 10. Minina E, Kreschel C, Naski MC, et al: Interaction of FGF, Ihh/ Pthlh, and BMP signaling integrates chondrocyte proliferation and hypertrophic differentiation. Dev Cell 3:439-449, 2002. 11. Lizarraga G, Lichtler A, Upholt WB, et al: Studies on the role of Cux1 in regulation of the onset of joint formation in the developing limb. Dev Biol 243:44-54, 2002. 12. Iwamoto M, Tamamura Y, Koyama E, et al: Transcription factor ERG and joint and articular cartilage formation during mouse limb and spine skeletogenesis. Dev Biol 305:40-51, 2007. 13. Hartmann C, Tabin CJ: Wnt-14 plays a pivotal role in inducing synovial joint formation in the developing appendicular skeleton. Cell 104:341-351, 2001. 14. Tsumaki N, Tanaka K, Arikawa-Hirasawa E, et al: Role of CDMP-1 in skeletal morphogenesis: Promotion of mesenchymal cell recruitment and chondrocyte differentiation. J Cell Biol 144:161-173, 1999. 15. Archer CW, Dowthwaite GP, Francis-West P: Development of synovial joints. Birth Defects Res C Embryo Today 69:144-155, 2003. 16. Storm EE, Kingsley DM: GDF5 coordinates bone and joint formation during digit development. Dev Biol 209:11-27, 1999. 17. von der Mark H, von der Mark K, Gay S: Study of differential collagen synthesis during development of the chick embryo by immunofluorescence, I: Preparation of collagen type I and type II specific antibodies and their application to early stages of the chick embryo. Dev Biol 48:237-249, 1976. 18. Craig FM, Bentley G, Archer CW: The spatial and temporal pattern of collagens I and II and keratan sulphate in the developing chick metatarsophalangeal joint. Development 99:383-391, 1987. 19. Morrison EH, Ferguson MW, Bayliss MT, et al: The development of articular cartilage, I: The spatial and temporal patterns of collagen types. J Anat 189(Pt 1):9-22, 1996. 20. Pitsillides AA, Archer CW, Prehm P, et al: Alterations in hyaluronan synthesis during developing joint cavitation. J Histochem Cytochem 43:263-273, 1995.   21. Murphy JM, Heinegard R, McIntosh A, et al: Distribution of cartilage molecules in the developing mouse joint. Matrix Biol 18:487497, 1999.   22. Mundlos S, Olsen BR: Heritable diseases of the skeleton, Part II: Molecular insights into skeletal development-matrix components and their homeostasis. FASEB J 11:227-233, 1997.   23. Kavanagh E, Ashhurst DE: Development and aging of the articular cartilage of the rabbit knee joint: Distribution of biglycan, decorin, and matrilin-1. J Histochem Cytochem 47:1603-1616, 1999.   24. Nalin AM, Greenlee TK Jr, Sandell LJ: Collagen gene expression during development of avian synovial joints: Transient expression of types II and XI collagen genes in the joint capsule. Dev Dyn 203:352-362, 1995.   25. Ito MM, Kida MY: Morphological and biochemical re-evaluation of the process of cavitation in the rat knee joint: Cellular and cell strata alterations in the interzone. J Anat 197(Pt 4):659-679, 2000.   26. Mitrovic D: Development of the diarthrodial joints in the rat embryo. Am J Anat 151:475-485, 1978.   27. Kimura S, Shiota K: Sequential changes of programmed cell death in developing fetal mouse limbs and its possible roles in limb morphogenesis. J Morphol 229:337-346, 1996.   28. Colnot CI, Helms JA: A molecular analysis of matrix remodeling and angiogenesis during long bone development. Mech Dev 100:245250, 2001.



17



  29. Roach HI, Clarke NM: Physiological cell death of chondrocytes in vivo is not confined to apoptosis: New observations on the mammalian growth plate. J Bone Joint Surg Br 82:601-613, 2000.   30. Edwards JCW, Wilkinson LS, Soothill P, et al: Matrix metalloproteinases in the formation of human synovial joint cavities. J Anat 188(Pt 2):355-360, 1996.   31. Edwards JCW, Wilkinson LS, Jones HM, et al: The formation of human synovial joint cavities: A possible role for hyaluronan and CD44 in altered interzone cohesion. J Anat 185(Pt 2):355-367, 1994.   32. Drachman DB, Sokoloff L: The role of movement in embryonic joint development. Dev Biol 14:401, 1966.   33. Persson M: The role of movements in the development of sutural and diarthrodial joints tested by long-term paralysis of chick embryos. J Anat 137(Pt 3):591-599, 1983.   34. Hall BK, Miyake T: All for one and one for all: Condensations and the initiation of skeletal development. Bioessays 22:138-147, 2000.   35. Capdevila J, Izpisua Belmonte JC: Patterning mechanisms controlling vertebrate limb development. Annu Rev Cell Dev Biol 17: 87-132, 2001.   36. Tuan RS: Biology of developmental and regenerative skeletogenesis. Clin Orthop 427(Suppl):S105-S117, 2004.   37. Fell HB: The histogenesis of cartilage and bone in the long bones of the embryonic fowl. J Morphol Physiol 40:417-459, 1925.   38. Sandell LJ, Nalin AM, Reife RA: Alternative splice form of type II procollagen mRNA (IIA) is predominant in skeletal precursors and non-cartilaginous tissues during early mouse development. Dev Dyn 199:129-140, 1994.   39. DeLise AM, Fischer L, Tuan RS: Cellular interactions and signaling in cartilage development. Osteoarthritis Cartilage 8:309-334, 2000.   40. Eames BF, de la Fuente L, Helms JA: Molecular ontogeny of the skeleton. Birth Defects Res C Embryo Today 69:93-101, 2003.   41. Zwilling E: Limb morphogenesis. Dev Biol 28:12-17, 1972.   42. Tickle C: Patterning systems—from one end of the limb to the other. Dev Cell 4:449-458, 2003.   43. Niswander L: Pattern formation: Old models out on a limb. Nat Rev Genet 4:133-143, 2003.   44. Kengaku M, Capdevila J, Rodriguez-Esteban C, et al: Distinct WNT pathways regulating AER formation and dorsoventral polarity in the chick limb bud. Science 280:1274-1277, 1998.   45. Johnson RL, Riddle RD, Tabin CJ: Mechanisms of limb patterning. Curr Opin Genet Dev 4:535-542, 1994.   46. Kmita M, Tarchini B, Zakany J, et al: Early developmental arrest of mammalian limbs lacking HoxA/HoxD gene function. Nature 435:1113-1116, 2005.   47. Riddle RD, Johnson RL, Laufer E, et al: Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 75:1401-1416, 1993.   48. Yang Y, Niswander L: Interaction between the signaling molecules WNT7a and SHH during vertebrate limb development: Dorsal signals regulate anteroposterior patterning. Cell 80:939-947, 1995.   49. Laufer E, Nelson CE, Johnson RL, et al: Sonic hedgehog and Fgf-4 act through a signaling cascade and feedback loop to integrate growth and patterning of the developing limb bud. Cell 79:993-1003, 1994.   50. Riddle RD, Tabin C: How limbs develop. Sci Am 280:74-79, 1999.   51. Bitgood MJ, McMahon AP: Hedgehog and Bmp genes are coexpressed at many diverse sites of cell-cell interaction in the mouse embryo. Dev Biol 172:126-138, 1995.   52. Roberts DJ, Johnson RL, Burke AC, et al: Sonic hedgehog is an endodermal signal inducing Bmp-4 and Hox genes during induction and regionalization of the chick hindgut. Development 121:31633174, 1995.   53. Liu A, Wang B, Niswander LA: Mouse intraflagellar transport proteins regulate both the activator and repressor functions of Gli transcription factors. Development 132:3103-3111, 2005.   54. Parr BA, McMahon AP: Dorsalizing signal Wnt-7a required for normal polarity of D-V and A-P axes of mouse limb. Nature 374:350353, 1995.   55. Irvine KD, Vogt TF: Dorsal-ventral signalling in limb development. Curr Opin Cell Biol 8:867-876, 1997.   56. Pizette S, Niswander L: BMPs are required at two steps of limb chondrogenesis: Formation of prechondrogenic condensations and their differentiation into chondrocytes. Dev Biol 219:237-249, 2000.



18



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| 



Biology of the Normal Joint



  57. Derynck R, Zhang YE: Smad-dependent and Smad-independent pathways in TGF-β family signalling. Nature 425:577-584, 2003.   58. Yoon BS, Lyons KM: Multiple functions of BMPs in chondrogenesis. J Cell Biochem 93:93-103, 2004.   59. Yoon BS, Ovchinnikov DA, Yoshii I, et al: Bmpr1a and Bmpr1b have overlapping functions and are essential for chondrogenesis in vivo. Proc Natl Acad Sci U S A 102:5062-5067, 2005.   60. Ng LJ, Wheatley S, Muscat GE, et al: SOX9 binds DNA, activates transcription, and coexpresses with type II collagen during chondrogenesis in the mouse. Dev Biol 183:108-121, 1997.   61. Lefebvre V, Huang W, Harley VR, et al: SOX9 is a potent activator of the chondrocyte-specific enhancer of the pro α1(II) collagen gene. Mol Cell Biol 17:2336-2346, 1997.   62. Lefebvre V, Li P, de Crombrugghe B: A new long form of Sox5 (L-Sox5), Sox6 and Sox9 are coexpressed in chondrogenesis and cooperatively activate the type II collagen gene. EMBO J 17: 5718-5733, 1998.   63. Smits P, Li P, Mandel J, et al: The transcription factors L-Sox5 and Sox6 are essential for cartilage formation. Dev Cell 1:277-290, 2001.   64. Ducy P, Zhang R, Geoffroy V, et al: Osf2/Cbfa1: A transcriptional activator of osteoblast differentiation. Cell 89:747-754, 1997.   65. Otto F, Thornell AP, Crompton T, et al: Cbfa1, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell 89:765-771, 1997.   66. Xu SC, Harris MA, Rubenstein JLR, et al: Bone morphogenetic protein-2 (BMP-2) signaling to the Col2a1 gene in chondroblasts requires the homeobox gene Dlx-2. DNA Cell Biol 20:359-365, 2001.   67. Itoh N, Ornitz DM: Evolution of the Fgf and Fgfr gene families. Trends Genet 20:563-569, 2004.   68. Ornitz DM: FGF signaling in the developing endochondral skeleton. Cytokine Growth Factor Rev 16:205-213, 2005.   69. Beier F: Cell-cycle control and the cartilage growth plate. J Cell Physiol 202:1-8, 2005.   70. Sahni M, Ambrosetti DC, Mansukhani A, et al: FGF signaling inhibits chondrocyte proliferation and regulates bone development through the STAT-1 pathway. Genes Dev 13:1361-1366, 1999.   71. Liu Z, Xu J, Colvin JS, et al: Coordination of chondrogenesis and osteogenesis by fibroblast growth factor 18. Genes Dev 16:859-869, 2002.   72. Ohbayashi N, Shibayama M, Kurotaki Y, et al: FGF18 is required for normal cell proliferation and differentiation during osteogenesis and chondrogenesis. Genes Dev 16:870-879, 2002.   73. Kronenberg HM: PTHrP and skeletal development. Ann N Y Acad Sci 1068:1-13, 2006.   74. McMahon AP, Ingham PW, Tabin CJ: Developmental roles and clinical significance of hedgehog signaling. Curr Top Dev Biol 53:1-114, 2003.   75. Vortkamp A, Lee K, Lanske B, et al: Regulation of rate of cartilage differentiation by Indian hedgehog and PTH-related protein. Science 273:613-622, 1996.   76. Lanske B, Karaplis AC, Lee K, et al: PTH/PTHrP receptor in early development and Indian hedgehog-regulated bone growth. Science 273:663-666, 1996.   77. Kobayashi T, Soegiarto DW, Yang Y, et al: Indian hedgehog stimulates periarticular chondrocyte differentiation to regulate growth plate length independently of PTHrP. J Clin Invest 115:1734-1742, 2005.   78. Ferguson CM, Miclau T, Hu D, et al: Common molecular pathways in skeletal morphogenesis and repair. Ann N Y Acad Sci 857:33-42, 1998.   79. Ballock RT, O’Keefe RJ: The biology of the growth plate. J Bone Joint Surg Am 85A:715-726, 2003.   80. Provot S, Schipani E: Molecular mechanisms of endochondral bone development. Biochem Biophys Res Commun 328:658-665, 2005.   81. St-Jacques B, Hammerschmidt M, McMahon AP: Indian hedgehog signaling regulates proliferation and differentiation of chondrocytes and is essential for bone formation. Genes Dev 13:2072-2086, 1999.   82. Enomoto H, Enomoto-Iwamoto M, Iwamoto M, et al: Cbfa1 is a positive regulatory factor in chondrocyte maturation. J Biol Chem 275:8695-8702, 2000.   83. Kim IS, Otto F, Zabel B, et al: Regulation of chondrocyte differentiation by Cbfa1. Mech Dev 80:159-170, 1999.



  84. Takeda S, Bonnamy JP, Owen MJ, et al: Continuous expression of Cbfa1 in nonhypertrophic chondrocytes uncovers its ability to induce hypertrophic chondrocyte differentiation and partially rescues Cbfa1-deficient mice. Genes Dev 15:467-481, 2001.   85. Colnot C: Cellular and molecular interactions regulating skeletogenesis. J Cell Biochem 95:688-697, 2005.   86. Leboy P, Grasso-Knight G, D’Angelo M, et al: Smad-Runx interactions during chondrocyte maturation. J Bone Joint Surg Am 83A(Suppl 1):S15-S22, 2001.   87. Zheng Q, Zhou G, Morello R, et al: Type X collagen gene regulation by Runx2 contributes directly to its hypertrophic chondrocyte-specific expression in vivo. J Cell Biol 162:833-842, 2003.   88. Komori T, Yagi H, Nomura S, et al: Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell 89:755-764, 1997.   89. Jimenez MJ, Balbin M, Lopez JM, et al: Collagenase 3 is a target of Cbfa1, a transcription factor of the runt gene family involved in bone formation. Mol Cell Biol 19:4431-4442, 1999.   90. Inada M, Yasui T, Nomura S, et al: Maturational disturbance of chondrocytes in Cbfa1-deficient mice. Dev Dyn 214:279-290, 1999.   91. Ortega N, Behonick DJ, Werb Z: Matrix remodeling during endochondral ossification. Trends Cell Biol 14:86-93, 2004.   92. Colnot C, Lu C, Hu D, et al: Distinguishing the contributions of the perichondrium, cartilage, and vascular endothelium to skeletal development. Dev Biol 269:55-69, 2004.   93. Inada M, Wang Y, Byrne MH, et al: Critical roles for collagenase3 (Mmp13) in development of growth plate cartilage and in endochondral ossification. Proc Natl Acad Sci U S A 101:17192-17197, 2004.   94. Stickens D, Behonick DJ, Ortega N, et al: Altered endochondral bone development in matrix metalloproteinase 13-deficient mice. Development 131:5883-5895, 2004.   95. Jacenko O, Chan D, Franklin A, et al: A dominant interference collagen X mutation disrupts hypertrophic chondrocyte pericellular matrix and glycosaminoglycan and proteoglycan distribution in transgenic mice. Am J Pathol 159:2257-2269, 2001.   96. Gress CJ, Jacenko O: Growth plate compressions and altered hematopoiesis in collagen X null mice. J Cell Biol 149:983-993, 2000.   97. Vu TH, Shipley JM, Bergers G, et al: MMP-9/gelatinase B is a key regulator of growth plate angiogenesis and apoptosis of hypertrophic chondrocytes. Cell 93:411-422, 1998.   98. Gerber HP, Vu TH, Ryan AM, et al: VEGF couples hypertrophic cartilage remodeling, ossification and angiogenesis during endochondral bone formation. Nat Med 5:623-628, 1999. 99. Maes C, Carmeliet P, Moermans K, et al: Impaired angiogenesis and endochondral bone formation in mice lacking the vascular endothelial growth factor isoforms VEGF164 and VEGF188. Mech Dev 111:61-73, 2002. 100. Maes C, Stockmans I, Moermans K, et al: Soluble VEGF isoforms are essential for establishing epiphyseal vascularization and regulating chondrocyte development and survival. J Clin Invest 113:188-199, 2004. 101. Zelzer E, Olsen BR: Multiple roles of vascular endothelial growth factor (VEGF) in skeletal development, growth, and repair. Curr Top Dev Biol 65:169-187, 2005. 102. Zhou Z, Apte SS, Soininen R, et al: Impaired endochondral ossification and angiogenesis in mice deficient in membrane-type matrix metalloproteinase I. Proc Natl Acad Sci U S A 97:4052-4057, 2000. 103. Merida-Velasco JA, Sanchez-Montesinos I, Espin-Ferra J, et al: Development of the human knee joint. Anat Rec 248:269-278, 1997. 104. Merida-Velasco JA, Sanchez-Montesinos I, Espin-Ferra J, et al: Development of the human knee joint ligaments. Anat Rec 248: 259-268, 1997. 105. Izumi S, Takeya M, Takagi K, et al: Ontogenetic development of synovial A cells in fetal and neonatal rat knee joints. Cell Tissue Res 262:1-8, 1990. 106. Edwards JCW: Fibroblast biology: Development and differentiation of synovial fibroblasts in arthritis. Arthritis Res 2:344-347, 2000. 107. Hukkanen M, Konttinen YT, Rees RG, et al: Distribution of nerve endings and sensory neuropeptides in rat synovium, meniscus and bone. Int J Tissue React 14:1-10, 1992. 108. Holmes G, Niswander L: Expression of slit-2 and slit-3 during chick development. Dev Dyn 222:301-307, 2001.



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| 



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109. Merida-Velasco JR, Rodriguez-Vazquez JF, Merida-Velasco JA, et al: Development of the human temporomandibular joint. Anat Rec 255:20-33, 1999. 110. Bradley SJ: An analysis of self-differentiation of chick limb buds in chorio-allantoic grafts. J Anat 107:479-490, 1970. 111. Roberts S, Evans H, Trivedi J, et al: Histology and pathology of the human intervertebral disc. J Bone Joint Surg Am 88(Suppl 2):10-14, 2006. 112. Eyre DR, Matsui Y, Wu JJ: Collagen polymorphisms of the intervertebral disc. Biochem Soc Trans 30:844-848, 2001. 113. Hayes AJ, Benjamin M, Ralphs JR: Extracellular matrix in development of the intervertebral disc. Matrix Biol 20:107-121, 2001. 114. McAlinden A, Zhu Y, Sandell LJ: Expression of type II procollagens during development of the human intervertebral disc. Biochem Soc Trans 30:831-838, 2001. 115. Zhu Y, McAlinden A, Sandell LJ: Type IIA procollagen in development of the human intervertebral disc: Regulated expression of the NH(2)-propeptide by enzymic processing reveals a unique developmental pathway. Dev Dyn 220:350-362, 2001. 116. Pizette S, Niswander L: Early steps in limb patterning and chondrogenesis. Novartis Found Symp 232:23-36; discussion 36-46, 2001. 117. Benjamin M, Ralphs JR: Tendons and ligaments—an overview. Histol Histopathol 12:1135-1144, 1997. 118. Poole AR: Cartilage in health and disease. In Koopman W (ed): Arthritis and Allied Conditions: A Textbook of Rheumatology. Philadelphia, Lippincott Williams & Wilkins, 2005, pp 223-269. 119. Henderson B, Pettipher ER: The synovial lining cell: Biology and pathobiology. Semin Arthritis Rheum 15:1-32, 1985. 120. Haywood L, Walsh DA: Vasculature of the normal and arthritic synovial joint. Histol Histopathol 16:277-284, 2001. 121. Tak PP, Bresnihan B: The pathogenesis and prevention of joint damage in rheumatoid arthritis: Advances from synovial biopsy and tissue analysis. Arthritis Rheum 43:2619-2633, 2000. 122. Firestein GS: Evolving concepts of rheumatoid arthritis. Nature 423:356-361, 2003. 123. Meyer LH, Franssen L, Pap T: The role of mesenchymal cells in the pathophysiology of inflammatory arthritis. Best Pract Res Clin Rheumatol 20:969-981, 2006. 124. Knedla A, Neumann E, Muller-Ladner U: Developments in the synovial biology field 2006. Arthritis Res Ther 9:209, 2007. 125. Szekanecz Z, Koch AE: Macrophages and their products in rheumatoid arthritis. Curr Opin Rheumatol 19:289-295, 2007. 126. Bassleer R, Lhosest-Gauthier M-P, Renard A-M, et al: Histological structure and functions of synovium. In Franchimont P (ed): Articular Synovium. Basel, Karger, 1982, pp 1-26. 127. Pasquali-Ronchetti I, Frizziero L, Guerra D, et al: Aging of the human synovium: An in vivo and ex vivo morphological study. Semin Arthritis Rheum 21:400-414, 1992. 128. Barland P, Novikoff AB, Hamerman D: Electron microscopy of the human synovial membrane. J Cell Biol 14:207-220, 1962. 129. Wilkinson LS, Pitsillides AA, Worrall JG, et al: Light microscopic characterization of the fibroblast-like synovial intimal cell (synoviocyte). Arthritis Rheum 35:1179-1184, 1992. 130. Smith MD, Barg E, Weedon H, et al: Microarchitecture and protective mechanisms in synovial tissue from clinically and arthroscopically normal knee joints. Ann Rheum Dis 62:303-307, 2003. 131. Naito M, Umeda S, Takahashi K, et al: Macrophage differentiation and granulomatous inflammation in osteopetrotic mice (op/op) defective in the production of CSF-1. Mol Reprod Dev 46:85-91, 1997. 132. Smith SC, Folefac VA, Osei DK, et al: An immunocytochemical study of the distribution of proline-4-hydroxylase in normal, osteoarthritic and rheumatoid arthritic synovium at both the light and electron microscopic level. Br J Rheumatol 37:287-291, 1998. 133. Qu Z, Garcia CH, O’Rourke LM, et al: Local proliferation of fibroblast-like synoviocytes contributes to synovial hyperplasia: Results of proliferating cell nuclear antigen/cyclin, c-myc, and nucleolar organizer region staining. Arthritis Rheum 37:212-220, 1994. 134. Walsh DA, Mapp PI, Wharton J, et al: Neuropeptide degrading enzymes in normal and inflamed human synovium. Am J Pathol 142:1610-1621, 1993. 135. Morales-Ducret J, Wayner E, Elices MJ, et al: α4/β1 integrin (VLA4) ligands in arthritis: Vascular cell adhesion molecule-1 expression in synovium and on fibroblast-like synoviocytes. J Immunol 149: 1424-1431, 1992.



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136. Demaziere A, Athanasou NA: Adhesion receptors of intimal and subintimal cells of the normal synovial membrane. J Pathol 168: 209-215, 1992. 137. Johnson BA, Haines GK, Harlow LA, et al: Adhesion molecule expression in human synovial tissue. Arthritis Rheum 36:137-146, 1993. 138. Valencia X, Higgins JM, Kiener HP, et al: Cadherin-11 provides specific cellular adhesion between fibroblast-like synoviocytes. J Exp Med 200:1673-1679, 2004. 139. Lee DM, Kiener HP, Agarwal SK, et al: Cadherin-11 in synovial lining formation and pathology in arthritis. Science 315:1006-1010, 2007. 140. Simkin PA: Physiology of normal and abnormal synovium. Semin Arthritis Rheum 21:179-183, 1991. 141. Firestein GS: Starving the synovium: Angiogenesis and inflammation in rheumatoid arthritis. J Clin Invest 103:3-4, 1999. 142. Walsh DA: Angiogenesis and arthritis. Rheumatology (Oxf) 38: 103-112, 1999. 143. Walsh DA, Wade M, Mapp PI, et al: Focally regulated endothelial proliferation and cell death in human synovium. Am J Pathol 152:691-702, 1998. 144. Koch AE: Review: Angiogenesis: Implications for rheumatoid arthritis. Arthritis Rheum 41:951-962, 1998. 145. Storgard CM, Stupack DG, Jonczyk A, et al: Decreased angiogenesis and arthritic disease in rabbits treated with an αvβ3 antagonist. J Clin Invest 103:47-54, 1999. 146. Uchida T, Nakashima M, Hirota Y, et al: Immunohistochemical localisation of protein tyrosine kinase receptors Tie-1 and Tie-2 in synovial tissue of rheumatoid arthritis: Correlation with angiogenesis and synovial proliferation. Ann Rheum Dis 59:607-614, 2000. 147. Gravallese EM, Pettit AR, Lee R, et al: Angiopoietin-1 is expressed in the synovium of patients with rheumatoid arthritis and is induced by tumour necrosis factor α. Ann Rheum Dis 62:100-107, 2003. 148. Mapp PI: Innervation of the synovium. Ann Rheum Dis 54:398-403, 1995. 149. Seegers HC, Hood VC, Kidd BL, et al: Enhancement of angiogenesis by endogenous substance P release and neurokinin-1 receptors during neurogenic inflammation. J Pharmacol Exp Ther 306:8-12, 2003. 150. Walsh DA, Catravas J, Wharton J: Angiotensin converting enzyme in human synovium: Increased stromal [(125)I]351A binding in rheumatoid arthritis. Ann Rheum Dis 59:125-131, 2000. 151. Mapp PI, Klocke R, Walsh DA, et al: Localization of 3-­nitrotyrosine to rheumatoid and normal synovium. Arthritis Rheum 44:15341539, 2001. 152. Pablos JL, Santiago B, Galindo M, et al: Synoviocyte-derived CXCL12 is displayed on endothelium and induces angiogenesis in rheumatoid arthritis. J Immunol 170:2147-2152, 2003. 153. Ferrell WR, Crighton A, Sturrock RD: Position sense at the proximal interphalangeal joint is distorted in patients with rheumatoid arthritis of finger joints. Exp Physiol 77:675-680, 1992. 154. Dee R: Structure and function of hip joint innervation. Ann R Coll Surg Engl 45:357-374, 1969. 155. Buma P, Verschuren C, Versleyen D, et al: Calcitonin gene-related peptide, substance P and GAP-43/B-50 immunoreactivity in the normal and arthrotic knee joint of the mouse. Histochemistry 98: 327-339, 1992. 156. Hukkanen M, Platts LA, Corbett SA, et al: Reciprocal age-related changes in GAP-43/B-50, substance P and calcitonin gene-related peptide (CGRP) expression in rat primary sensory neurones and their terminals in the dorsal horn of the spinal cord and subintima of the knee synovium. Neurosci Res 42:251-260, 2002. 157. McDougall JJ, Watkins L, Li Z: Vasoactive intestinal peptide (VIP) is a modulator of joint pain in a rat model of osteoarthritis. Pain 123:98-105, 2006. 158. Niissalo S, Hukkanen M, Imai S, et al: Neuropeptides in experimental and degenerative arthritis. Ann N Y Acad Sci 966:384-399, 2002. 159. Saito T, Koshino T: Distribution of neuropeptides in synovium of the knee with osteoarthritis. Clin Orthop 376:172-182, 2000. 160. New HV, Mudge AW: Calcitonin gene-related peptide regulates muscle acetylcholine receptor synthesis. Nature 323:809-811, 1986. 161. Schaible HG, Ebersberger A, Von Banchet GS: Mechanisms of pain in arthritis. Ann N Y Acad Sci 966:343-354, 2002. 162. McDougall JJ: Arthritis and pain: Neurogenic origin of joint pain. Arthritis Res Ther 8:220, 2006. 163. Coggeshall RE, Hong KA, Langford LA, et al: Discharge characteristics of fine medial articular afferents at rest and during passive movements of inflamed knee joints. Brain Res 272:185-188, 1983.



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164. Benjamin M, Ralphs JR: The cell and developmental biology of tendons and ligaments. Int Rev Cytol 196:85-130, 2000. 165. Wang JH: Mechanobiology of tendon. J Biomech 39:1563-1582, 2006. 166. Vogel KG, Peters JA: Histochemistry defines a proteoglycan-rich layer in bovine flexor tendon subjected to bending. J Musculoskelet Neuronal Interact 5:64-69, 2005. 167. Knott L, Tarlton JF, Bailey AJ: Chemistry of collagen cross-linking: Biochemical changes in collagen during the partial mineralization of turkey leg tendon. Biochem J 322(Pt 2):535-542, 1997. 168. Ng GY, Oakes BW, Deacon OW, et al: Long-term study of the biochemistry and biomechanics of anterior cruciate ligament-patellar tendon autografts in goats. J Orthop Res 14:851-856, 1996. 169. Canty EG, Starborg T, Lu Y, et al: Actin filaments are required for fibripositor-mediated collagen fibril alignment in tendon. J Biol Chem 281:38592-38598, 2006. 170. Richardson SH, Starborg T, Lu Y, et al: Tendon development requires regulation of cell condensation and cell shape via cadherin-11­mediated cell-cell junctions. Mol Cell Biol 27:6218-6228, 2007. 171. Kannus P, Jozsa L, Kvist M, et al: The effect of immobilization on myotendinous junction: An ultrastructural, histochemical and immunohistochemical study. Acta Physiol Scand 144:387-394, 1992. 172. Tan AL, Toumi H, Benjamin M, et al: Combined high-resolution magnetic resonance imaging and histological examination to explore the role of ligaments and tendons in the phenotypic expression of early hand osteoarthritis. Ann Rheum Dis 65:1267-1272, 2006. 173. Dalton S, Cawston TE, Riley GP, et al: Human shoulder tendon biopsy samples in organ culture produce procollagenase and tissue inhibitor of metalloproteinases. Ann Rheum Dis 54:571-577, 1995. 174. Jain A, Nanchahal J, Troeberg L, et al: Production of cytokines, vascular endothelial growth factor, matrix metalloproteinases, and tissue inhibitor of metalloproteinases 1 by tenosynovium demonstrates its potential for tendon destruction in rheumatoid arthritis. Arthritis Rheum 44:1754-1760, 2001. 175. Tillander B, Franzen L, Norlin R: Fibronectin, MMP-1 and histologic changes in rotator cuff disease. J Orthop Res 20:1358-1364, 2002. 176. Jarvinen TA, Jozsa L, Kannus P, et al: Mechanical loading regulates the expression of tenascin-C in the myotendinous junction and tendon but does not induce de novo synthesis in the skeletal muscle. J Cell Sci 116:857-866, 2003. 177. Berenson MC, Blevins FT, Plaas AH, et al: Proteoglycans of human rotator cuff tendons. J Orthop Res 14:518-525, 1996. 178. Waggett AD, Ralphs JR, Kwan AP, et al: Characterization of collagens and proteoglycans at the insertion of the human Achilles tendon. Matrix Biol 16:457-470, 1998. 179. Fukuta S, Oyama M, Kavalkovich K, et al: Identification of types II, IX and X collagens at the insertion site of the bovine achilles tendon. Matrix Biol 17:65-73, 1998. 180. Vogel KG, Meyers AB: Proteins in the tensile region of adult bovine deep flexor tendon. Clin Orthop 367:S344-S355, 1999. 181. Thomopoulos S, Marquez JP, Weinberger B, et al: Collagen fiber orientation at the tendon to bone insertion and its influence on stress concentrations. J Biomech 39:1842-1851, 2006. 182. Rodeo SA, Kawamura S, Kim HJ, et al: Tendon healing in a bone tunnel differs at the tunnel entrance versus the tunnel exit: An effect of graft-tunnel motion? Am J Sports Med 34:1790-1800, 2006. 183. Sharma P, Maffulli N: Biology of tendon injury: Healing, modeling and remodeling. J Musculoskelet Neuronal Interact 6:181-190, 2006. 184. Rees JD, Wilson AM, Wolman RL: Current concepts in the management of tendon disorders. Rheumatology (Oxf) 45:508-521, 2006. 185. Kumagai J, Sarkar K, Uhthoff HK: The collagen types in the attachment zone of rotator cuff tendons in the elderly: An immunohistochemical study. J Rheumatol 21:2096-2100, 1994. 186. Ameye L, Aria D, Jepsen K, et al: Abnormal collagen fibrils in tendons of biglycan/fibromodulin-deficient mice lead to gait impairment, ectopic ossification, and osteoarthritis. FASEB J 16:673-680, 2002. 187. Hoffmann A, Pelled G, Turgeman G, et al: Neotendon formation induced by manipulation of the Smad8 signalling pathway in mesenchymal stem cells. J Clin Invest 116:940-952, 2006. 188. Woo SL, Abramowitch SD, Kilger R, et al: Biomechanics of knee ligaments: Injury, healing, and repair. J Biomech 39:1-20, 2006. 189. Hoffmann A, Gross G: Tendon and ligament engineering: From cell biology to in vivo application. Regen Med 1:563-574, 2006.



190. Bland YS, Ashhurst DE: Changes in the distribution of fibrillar collagens in the collateral and cruciate ligaments of the rabbit knee joint during fetal and postnatal development. Histochem J 28:325-334, 1996. 191. Nawata K, Minamizaki T, Yamashita Y, et al: Development of the attachment zones in the rat anterior cruciate ligament: Changes in the distributions of proliferating cells and fibrillar collagens during postnatal growth. J Orthop Res 20:1339-1344, 2002. 192. Frank CB, Hart DA, Shrive NG: Molecular biology and biomechanics of normal and healing ligaments—a review. Osteoarthritis Cartilage 7:130-140, 1999. 193. Kaufmann P, Bose P, Prescher A: New insights into the soft-tissue anatomy anterior to the patella. Lancet 363:586, 2004. 194. Arnoczky SP, McDevitt CA: The meniscus: Structure, function repair, and replacement. In Buckwalter JL, Einhorn TA, Simon SR (eds): Orthopaedic Basic Science: Biology and Biomechanics of the Musculoskeletal System. Park Ridge, Ill, American Academy of Orthopaedic Surgeons, 2000, pp 531-546. 195. Sweigart MA, Athanasiou KA: Toward tissue engineering of the knee meniscus. Tissue Eng 7:111-129, 2001. 196. Fairbank T: Knee joint changes after meniscectomy. J Bone Joint Surg Br 30:664, 1948. 197. Murrell GA, Maddali S, Horovitz L, et al: The effects of time course after anterior cruciate ligament injury in correlation with meniscal and cartilage loss. Am J Sports Med 29:9-14, 2001. 198. Messner K, Gao J: The menisci of the knee joint: Anatomical and functional characteristics, and a rationale for clinical treatment. J Anat 193(Pt 2):161-178, 1998. 199. Mine T, Kimura M, Sakka A, et al: Innervation of nociceptors in the menisci of the knee joint: An immunohistochemical study. Arch Orthop Trauma Surg 120:201-204, 2000. 200. Arnoczky SP, Warren RF: The microvasculature of the meniscus and its response to injury: An experimental study in the dog. Am J Sports Med 11:131-141, 1983. 201. Eyre DR, Muir H: The distribution of different molecular species of collagen in fibrous, elastic and hyaline cartilages of the pig. Biochem J 151:595-602, 1975. 202. McDevitt CA, Mukherjee S, Kambic H, et al: Emerging concepts of the cell biology of the meniscus. Curr Opin Orthop 13:345-350, 2002. 203. Petersen W, Tillmann B: Collagenous fibril texture of the human knee joint menisci. Anat Embryol (Berl) 197:317-324, 1998. 204. Fithian DC, Kelly MA, Mow VC: Material properties and structure-function relationships in the menisci. Clin Orthop 252:19-31, 1990. 205. Roughley PJ, White RJ: The dermatan sulfate proteoglycans of the adult human meniscus. J Orthop Res 10:631-637, 1992. 206. Bland YS, Ashhurst DE: Changes in the content of the fibrillar collagens and the expression of their mRNAs in the menisci of the rabbit knee joint during development and ageing. Histochem J 28:265-274, 1996. 207. McAlinden A, Dudhia J, Bolton MC, et al: Age-related changes in the synthesis and mRNA expression of decorin and aggrecan in human meniscus and articular cartilage. Osteoarthritis Cartilage 9:33-41, 2001. 208. Ghadially FN, Lalonde JM, Wedge JH: Ultrastructure of normal and torn menisci of the human knee joint. J Anat 136(Pt 4):773-791, 1983. 209. Hellio Le Graverand MP, Ou Y, Schield-Yee T, et al: The cells of the rabbit meniscus: Their arrangement, interrelationship, morphological variations and cytoarchitecture. J Anat 198:525-535, 2001. 210. Kambic HE, Futani H, McDevitt CA: Cell, matrix changes and α-smooth muscle actin expression in repair of the canine meniscus. Wound Repair Regen 8:554-561, 2000. 211. Ahluwalia S, Fehm M, Murray MM, et al: Distribution of smooth muscle actin-containing cells in the human meniscus. J Orthop Res 19:659-664, 2001. 212. Goldring MB: The muskuloskeletal system, B: Articular cartilage. In Klippel JH, Crofford LJ, Stone JH, et al (eds): Primer on the Rheumatic Diseases. Atlanta, Arthritis Foundation, 2001, pp 10-16. 213. Benedek TG: A history of the understanding of cartilage. Osteoarthritis Cartilage 14:203-209, 2006.



PART 1 



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STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



214. Bailey AJ, Mansell JP, Sims TJ, et al: Biochemical and mechanical properties of subchondral bone in osteoarthritis. Biorheology 41: 349-358, 2004. 215. Green WT Jr, Martin GN, Eanes ED, et al: Microradiographic study of the calcified layer of articular cartilage. Arch Pathol 90:151-158, 1970. 216. Reimann I, Mankin HJ, Trahan C: Quantitative histologic analyses of articular cartilage and subchondral bone from osteoarthritic and normal human hips. Acta Orthop Scand 48:63-73, 1977. 217. Radin EL, Rose RM: Role of subchondral bone in the initiation and progression of cartilage damage. Clin Orthop 213:34-40, 1986. 218. Burr DB: Anatomy and physiology of the mineralized tissues: Role in the pathogenesis of osteoarthrosis. Osteoarthritis Cartilage 12(Suppl A): S20-S30, 2004. 219. Buckland-Wright C: Subchondral bone changes in hand and knee osteoarthritis detected by radiography. Osteoarthritis Cartilage 12(Suppl A):S10-S19, 2004. 220. Mrosek EH, Lahm A, Erggelet C, et al: Subchondral bone trauma causes cartilage matrix degeneration: An immunohistochemical analysis in a canine model. Osteoarthritis Cartilage 14:171-178, 2006. 221. Lane LB, Villacin A, Bullough PG: The vascularity and remodelling of subchondrial bone and calcified cartilage in adult human femoral and humeral heads: An age- and stress-related phenomenon. J Bone Joint Surg Br 59:272-278, 1977. 222. Walsh DA, Bonnet CS, Turner EL, et al: Angiogenesis in the synovium and at the osteochondral junction in osteoarthritis. Osteoarthritis Cartilage 15:743-751, 2007. 223. Bullough PG: The role of joint architecture in the etiology of arthritis. Osteoarthritis Cartilage 12(Suppl A):S2-S9, 2004. 224. Messent EA, Ward RJ, Tonkin CJ, et al: Differences in trabecular structure between knees with and without osteoarthritis quantified by macro and standard radiography, respectively. Osteoarthritis Cartilage 14:1302-1305, 2006. 225. Coats AM, Zioupos P, Aspden RM: Material properties of subchondral bone from patients with osteoporosis or osteoarthritis by microindentation testing and electron probe microanalysis. Calcif Tissue Int 73:66-71, 2003. 226. El Hajjaji H, Williams JM, Devogelaer JP, et al: Treatment with calcitonin prevents the net loss of collagen, hyaluronan and proteoglycan aggregates from cartilage in the early stages of canine experimental osteoarthritis. Osteoarthritis Cartilage 12:904-911, 2004. 227. Bagger YZ, Tanko LB, Alexandersen P, et al: Oral salmon calcitonin induced suppression of urinary collagen type II degradation in postmenopausal women: A new potential treatment of osteoarthritis. Bone 37:425-430, 2005. 228. Spector TD: Bisphosphonates: Potential therapeutic agents for disease modification in osteoarthritis. Aging Clin Exp Res 15:413-418, 2003. 229. Ham KD, Carlson CS: Effects of estrogen replacement therapy on bone turnover in subchondral bone and epiphyseal metaphyseal cancellous bone of ovariectomized cynomolgus monkeys. J Bone Miner Res 19:823-829, 2004. 230. Komuro H, Olee T, Kuhn K, et al: The osteoprotegerin/receptor activator of nuclear factor ĸB/receptor activator of nuclear factor ĸB ligand system in cartilage. Arthritis Rheum 44:2768-2776, 2001. 231. Pettit AR, Ji H, von Stechow D, et al: TRANCE/RANKL knockout mice are protected from bone erosion in a serum transfer model of arthritis. Am J Pathol 159:1689-1699, 2001. 232. Zwerina J, Hayer S, Redlich K, et al: Activation of p38 MAPK is a key step in tumor necrosis factor-mediated inflammatory bone destruction. Arthritis Rheum 54:463-472, 2006. 233. Simkin PA, Bassett JE, Koh EM: Synovial perfusion in the human knee: A methodologic analysis. Semin Arthritis Rheum 25:56-66, 1995. 234. Kushner I, Somerville JA: Permeability of human synovial membrane to plasma proteins: Relationship to molecular size and inflammation. Arthritis Rheum 14:560-570, 1971. 235. Simkin PA: Fluid dynamics of the joint space and trafficking of matrix products. In Seibel MJ, Robins SP, Bilezekian JP (eds): Dynamics of Bone and Cartilage Metabolism. New York, Academic Press, 2006, pp 451-456.



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236. Levick JR: A method for estimating macromolecular reflection by human synovium, using measurements of intra-articular half lives. Ann Rheum Dis 57:339-344, 1998. 237. Levick JR, McDonald JN: Fluid movement across synovium in healthy joints: Role of synovial fluid macromolecules. Ann Rheum Dis 54:417-423, 1995. 238. Myers SL, Brandt KD: Effects of synovial fluid hyaluronan concentration and molecular size on clearance of protein from the canine knee. J Rheumatol 22:1732-1739, 1995. 239. Gaffney K, Williams RB, Jolliffe VA, et al: Intra-articular pressure changes in rheumatoid and normal peripheral joints. Ann Rheum Dis 54:670-673, 1995. 240. Wallis WJ, Simkin PA, Nelp WB: Protein traffic in human synovial effusions. Arthritis Rheum 30:57-63, 1987. 241. Myers SL, O’Connor BL, Brandt KD: Accelerated clearance of albumin from the osteoarthritic knee: Implications for interpretation of concentrations of “cartilage markers” in synovial fluid. J Rheumatol 23:1744-1748, 1996. 242. Hlavacek M: The role of synovial fluid filtration by cartilage in lubrication of synovial joints, II: Squeeze-film lubrication: Homogeneous filtration. J Biomech 26:1151-1160, 1993. 243. Jin ZM, Dowson D, Fisher J: The effect of porosity of articular cartilage on the lubrication of a normal human hip joint. Proc Inst Mech Eng [H] 206:117-124, 1992. 244. Swann DA, Silver FH, Slayter HS, et al: The molecular structure and lubricating activity of lubricin isolated from bovine and human synovial fluids. Biochem J 225:195-201, 1985. 245. Jay GD, Tantravahi U, Britt DE, et al: Homology of lubricin and superficial zone protein (SZP): Products of megakaryocyte stimulating factor (MSF) gene expression by human synovial fibroblasts and articular chondrocytes localized to chromosome 1q25. J Orthop Res 19:677-687, 2001. 246. Flannery CR, Hughes CE, Schumacher BL, et al: Articular cartilage superficial zone protein (SZP) is homologous to megakaryocyte stimulating factor precursor and is a multifunctional proteoglycan with potential growth-promoting, cytoprotective, and lubricating properties in cartilage metabolism. Biochem Biophys Res Commun 254:535-541, 1999. 247. Ikegawa S, Sano M, Koshizuka Y, et al: Isolation, characterization and mapping of the mouse and human PRG4 (proteoglycan 4) genes. Cytogenet Cell Genet 90:291-297, 2000. 248. Schmidt TA, Schumacher BL, Klein TJ, et al: Synthesis of proteoglycan 4 by chondrocyte subpopulations in cartilage explants, monolayer cultures, and resurfaced cartilage cultures. Arthritis Rheum 50:2849-2857, 2004. 249. Jay GD, Lane BP, Sokoloff L: Characterization of a bovine synovial fluid lubricating factor, III: The interaction with hyaluronic acid. Connect Tissue Res 28:245-255, 1992. 250. Williams PF 3rd, Powell GL, LaBerge M: Sliding friction analysis of phosphatidylcholine as a boundary lubricant for articular cartilage. Proc Inst Mech Eng [H] 207:59-66, 1993. 251. Hills BA: Boundary lubrication in vivo. Proc Inst Mech Eng 214: 83-94, 2000. 252. Jay GD, Harris DA, Cha CJ: Boundary lubrication by lubricin is mediated by O-linked β(1-3)Gal-GalNAc oligosaccharides. Glycoconj J 18:807-815, 2001. 253. Stockwell RA: Lipid content of human costal and articular cartilage. Ann Rheum Dis 26:481-486, 1967. 254. Pickard JE, Fisher J, Ingham E, et al: Investigation into the effects of proteins and lipids on the frictional properties of articular cartilage. Biomaterials 19:1807-1812, 1998. 255. Hunter W: On the structure and diseases of articulating cartilage. Philos Trans R Soc Lond Biol 42:514, 1743. 256. Brem H, Folkman J: Inhibition of tumor angiogenesis mediated by cartilage. J Exp Med 141:427-439, 1975. 257. Kuettner KE, Pauli BU: Inhibition of neovascularization by a cartilage factor. Ciba Found Symp 100:163-173, 1983. 258. Moses MA, Sudhalter J, Langer R: Identification of an inhibitor of neovascularization from cartilage. Science 248:1408-1410, 1990. 259. Pita JC, Howell DS: Micro-biochemical studies of cartilage. In Sokoloff L (ed): The Joints and Synovial Fluid. New York, Academic Press, 1978, p 273. 260. Bywaters E: The metabolism of joint tissues. J Pathol Bacteriol 44:247, 1937.



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261. Naughton DP, Haywood R, Blake DR, et al: A comparative evaluation of the metabolic profiles of normal and inflammatory knee-joint synovial fluids by high resolution proton NMR spectroscopy. FEBS Lett 332:221-225, 1993. 262. Strangeways T: The nutrition of the articular cartilage. BMJ 1: 661, 1920. 263. Maroudas A, Bullough P, Swanson SA, et al: The permeability of articular cartilage. J Bone Joint Surg Br 50:166-177, 1968. 264. Hadler NM: Synovial fluids facilitate small solute diffusivity. Ann Rheum Dis 39:580-585, 1980. 265. O’Hara BP, Urban JP, Maroudas A: Influence of cyclic loading on the nutrition of articular cartilage. Ann Rheum Dis 49:536-539, 1990. 266. Lewis P, McCutchen CW: Experimental evidence for weeping lubrication in mammalian joints. Nature 184:1285, 1959.



267. Mital MA, Millington PF: Osseous pathway of nutrition to articular cartilage of the human femoral head. Lancet 1:842, 1970. 268. Bromley M, Bertfield H, Evanson JM, et al: Bidirectional erosion of cartilage in the rheumatoid knee joint. Ann Rheum Dis 44:676-681, 1985. 269. Lane LB, Bullough PG: Age-related changes in the thickness of the calcified zone and the number of tidemarks in adult human articular cartilage. J Bone Joint Surg Br 62:372-375, 1980. 270. Neusel E, Graf J: The influence of subchondral vascularisation on chondromalacia patellae. Arch Orthop Trauma Surg 115:313-315, 1996.



2



Synovium BARRY BRESNIHAN  •  ADRIENNE M. FLANAGAN



KEY POINTS The synovium provides nutrients to cartilage and produces lubricants for the joint. The intimal lining of the synovium introduces macrophagelike and fibroblast-like synoviocytes. The sublining contains scattered immune cells, fibroblasts, blood vessels, and fat cells. Fibroblast-like synoviocytes in the intimal lining express specialized proteins that synthesize proteoglycans such as hyaluronic acid.



STRUCTURE The synovium is a membranous structure that extends from the margins of articular cartilage and lines the capsule of diar­ throdial joints, including the temporomandibular joint1 and the facet joints of vertebral bodies (Fig. 2-1).2 The healthy synovium covers intra-articular tendons and ­ ligaments, and fat pads, but not articular cartilage or meniscal tissue. Synovium also ensheathes tendons where they pass beneath ligamentous bands. Normally, the synovial membrane has two components—the intima, or lining cells, and the sub­ intima, otherwise referred to as the sublining or supportive layer. The intima represents the interface between the cav­ ity containing synovial fluid and the subintimal layer. There is no well-formed basement membrane to separate the intima from the subintima. The subintima is composed of fibrovascular connective tissue and merges with the densely collagenous fibrous joint capsule. SYNOVIAL LINING CELLS The synovial intimal layer is composed of synovial lining cells (SLCs), which have an epithelial-like arrangement on the luminal aspect of the joint cavity. SLCs, termed synoviocytes, are one to three cells deep, depending on the anatomic location, and they extend 20 to 40 μm beneath the lining layer surface. The major and minor axes of SLCs measure 8 to 12 μm (major axis) and 6 to 8 μm (minor axis). SLCs have poorly defined cell borders and elliptical nuclei with generally a single small nucleolus.3 Ultrastructure of Synovial Lining Cells Transmission electron microscopic analysis shows that the intimal cells form a discontinuous layer, something not appre­ ciated under transmission light microscopy, so that the subin­ timal matrix is in direct contact with the synovial fluid (Fig. 2-2). The existence of two distinct cell types, type A and type B



SLCs, originally was described by Barland and associates,4 and several lines of evidence, including animal models, detailed ultrastructural studies, and immunohistochemical analysis, indicate that these cells represent macrophages (type A SLCs) and fibroblasts (type B SLCs). Studies of the SLC populations in a variety of species, including humans, have found that macrophages make up approximately 20% and fibroblast-like cells approximately 80% of the lining cell.5,6 The existence of the two cell types has been substantiated by similar findings in a wide variety of species, including hamsters, cats, dogs, guinea pigs, rabbits, mice, rats, and horses.6-14 Distinguishing the different cell populations that form the synovial lining is impossible by hematoxylin and eosin staining under transmission light microscopy. At an ultra­ structural level, the type A cells are characterized by a con­ spicuous Golgi apparatus, large vacuoles, and small vesicles, and contain little rough endoplasmic reticulum, giving them a macrophage-like phenotype (Fig. 2-3A and B). The plasma membrane of type A cells possesses numerous fine extensions, termed filopodia, which are characteristic of mac­ rophages. These cells are located for the most part on the lin­ ing surface, where it is more than one cell thick. Type A cells cluster at the tips of the synovial villi, and this uneven dis­ tribution at least partly explains early reports that suggested type A cells were the predominant intimal cell type.4,8 Type B SLCs have prominent cytoplasmic extensions that extend onto the surface of the synovial lining (Fig. 2-3C and D).15 Frequent invaginations are seen along the plasma membrane, and a large indented nucleus relative to the area of the surrounding cytoplasm is also a feature. Type B cells have abundant rough endoplasmic reticulum widely distrib­ uted in the cytoplasm, and the Golgi apparatus, vacuoles, and vesicles are generally inconspicuous, although some cells have small numbers of prominent vacuoles at their apical aspect. Type B SLCs also are known to contain lon­ gitudinal bundles of different-sized filaments, supporting their classification as fibroblasts. Desmosomes and gaplike junctions have been described in rat, mouse, and rabbit synovium, but the existence of these structures has never been documented in human SLCs. Cells exhibiting the ultrastructure of type A and type B SLCs have been classified as intermediate, or type AB. The existence of intermediate cells has been refuted on the basis of detailed electron microscopic studies, and it is now accepted that a proportion of type B cells have conspicu­ ous vacuoles, and that rough endoplasmic reticulum appears in activated macrophages.16,17 The putative existence of an intermediate SLC implies that type A and type B SLCs are part of the same cell lineage. This concept is contrary to all current evidence, which finds that type A and type B SLCs are histogenetically and functionally distinct. 23



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Figure 2-2  Transmission electron photomicrograph of synovial intimal cells. The cell on the left exhibits the dendritic appearance of a synovial intimal fibroblast (type B cell). Other overlying fibroblast dendrites can be observed. The presence of intercellular gaps allows the synovial fluid to be in direct contact with the synovial matrix. 500 µm Figure 2-1  The cartilage-synovium junction. Hyaline articular cartilage occupies the left half of this image, and fibrous capsule and synovial membrane occupy the right half. A sparse intimal lining layer with a fibrous subintima can be observed extending from the margin of the cartilage across the capsular surface to assume a more cellular intimal morphology with areolar subintima.



Immunohistochemical Profile of Synovial Intimal Cells Synovial Intimal Macrophages. Synovial macrophages and fibroblasts express lineage-specific molecules, which can be detected by immunohistochemistry. Synovial macrophages express common hematopoietic antigen CD45 (Fig. 2-4A); monocyte/macrophage receptors CD163 and CD97; and ly­ sosomal enzymes CD68 (Fig. 2-4B), neuron-specific esterase, and cathepsin B, L, and D. Cells expressing CD14, a mol­ ecule that acts as a coreceptor for the detection of bacterial lipopolysaccharide, and expressed by circulating monocytes and monocytes newly recruited to tissue, are rarely seen in the healthy intimal layer, but small numbers are found close to venules in the subintima.18-24 The Fcγ receptor, FcγRIII (CD16), expressed by Kupffer cells of the liver and type II alveolar macrophages of the lung, also is expressed on a subpopulation of synovial mac­ rophages.25-27 The synovial macrophage population also expresses the major histocompatibility complex (MHC) class II molecule which plays an important role in the immune response. More recently, the macrophages, which are responsible for the removal of debris, blood, and par­ ticulate material from the joint cavity and possess antigen processing properties, have been found to express a new complement-related protein, Z39Ig, a cell surface receptor and immunoglobulin superfamily member, which is involved in the induction of HLA-DR, and implicated in the reg­ ulation of phagocytosis and antigen-­mediated immune responses.28-30 The expression of the β2 integrin chains, CD18, CD11a, CD11b, and CD11c, varies; CD11a and CD11c may be absent, or weakly expressed, on a few lining cells.31,32 Osteoclasts, which are tartrate-resistant, acid phosphatase– positive, and express the αvβ3 vitronectin and calcitonin receptors, do not appear in the normal synovium. Synovial Intimal Fibroblasts. Synovial intimal and subin­ timal fibroblasts are indistinguishable by light microscopy. They are generally considered to be closely related in terms



of cell lineage, but because of their different microenviron­ ments, they do not always share the same phenotype. They possess prominent synthetic capacity and produce the es­ sential joint lubricants hyaluronic acid (HA) and lubricin.33 Intimal fibroblasts express uridine diphosphoglucose dehy­ drogenase (UDPGD), an enzyme involved in HA synthesis, which is recognized as a specific marker for this cell type. UDPGD converts UDP-glucose to UDP-glucuronate, one of the two substrates required by HA synthase for assembly of the HA polymer.34 CD44 expression, the nonintegrin re­ ceptor for HA, is expressed by all SLCs.32,35,36 Synovial fibroblasts also synthesize normal matrix com­ ponents, including fibronectin, laminin, collagens, pro­ teoglycans, lubricin, and other identified and unidentified proteins. They also have the capacity to produce large amounts of metalloproteinases, metalloproteinase inhibi­ tors, prostaglandins, and cytokines. This capacity must provide essential biologic advantages, but the complex physiologic mechanisms relevant to normal function are incompletely delineated. The expression of selected adhe­ sion molecules on synovial fibroblasts probably facilitates the trafficking of some cell populations, such as neutrophils, into the synovial fluid, and the retention of others, such as mononuclear leukocytes, in the synovial tissue. Metal­ loproteinases, cytokines, adhesion molecules, and other cell surface molecules are strikingly upregulated in inflamma­ tory states. Specialized intimal fibroblasts also express many other molecules that are not expressed by the intimal macrophage population, including decay-accelerating factor (CD55), previously identified by the antibody Mab67; vascular cell adhesion molecule 1; intracellular adhesion molecule33,37-40; and cadherin 11.41,42 PGP.95, a neuronal marker, is reported as being specific for type B synoviocytes in horses.43 Decayaccelerating factor, also expressed on the cells of other body cavities and cells in bone marrow, interacts with CD97, a glycoprotein that is present on the surface of most acti­ vated leukocytes, including intimal macrophages, and is thought to be involved in the signaling processes early after leukocyte activation.44,45 In contrast, FcγRIII is expressed only by macrophages when they are in close contact with decay-accelerating factor–positive fibroblasts, or decayaccelerating factor–coated fibrillin-1 microfibrils in the extracellular matrix.26 Cadherins are a class of tissue-restricted transmem­ brane proteins that play important roles in homophilic



PART 1 



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STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



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Figure 2-3  Transmission electron photomicrographs of synovial intimal macrophages (type A cells) and fibroblasts (type B cells). A, Low-powered magnification showing the surface fine filopodia, characteristic of macrophages, and a smooth-surfaced nucleus. B, The boxed area in A is shown at a higher magnification and reveals numerous vesicles characteristic of macrophages. The absence of rough endoplasmic reticulum also is noted. C, The convoluted nucleus along with the prominent rough endoplasmic reticulum (boxed area) is characteristic of a synovial intimal fibroblast (type B cell).  D, The rough endoplasmic reticulum is shown at greater magnification.



intercellular adhesion and are involved in maintaining the integrity of tissue architecture. Cadherin 11, which was cloned from rheumatoid arthritis synovial tissue, also is expressed in normal synovial intimal fibroblasts, but not in intimal macrophages. The finding that fibroblasts trans­ fected with cadherin 11 are induced to form a lining-like structure in vitro implicates this molecule in the architec­ tural organization of the synovial lining.41,42,46 This sugges­ tion is supported by the observation that cadherin-deficient mice have a hypoplastic synovial lining and are resistant to inflammatory arthritis.47



β1 and β3 integrins are present on all SLCs, forming recep­ tors for laminin (CD49f and CD49b), collagen types I and IV (CD49b), vitronectin (CD51), and fibronectin (CD49d and CD49e). In contrast, the integrin collagen receptors, CD49a, CD54 (a member of the immunoglobulin superfam­ ily), and CD4 and CD62 (selectin) present on lymphocytes, and involved in their homing to high endothelial venules, are not observed on these cells. CD31 (platelet–endothelial cell adhesion molecule), a member of the immunoglobulin superfamily that is expressed on endothelial cells, platelets, and monocytes, is only weakly expressed on SLCs.32



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Figure 2-4  Transmitted light photomicrographs depicting synovial intimal macrophages by immunohistochemistry. A and B, Macrophages are decorated with CD45 (arrow in A) and CD68 (B), markers that identify hematopoietic cells (CD45) and macrophages (CD68).



Turnover of Synovial Lining Cells Proliferation of SLCs in humans is low, as shown when normal human synovial explants, exposed to a pulse of 3H thymidine, resulted in the SLCs having a labeling index of approximately 0.05% to 0.3%48; this bears a striking con­ trast with labeling indices of approximately 50% for bowel crypt epithelium. Similar evidence of low proliferation has been found in the synovium of rats and rabbits. The advent of immunohistochemistry saw this observation substanti­ ated when Revell and others reported that the proportion of SLCs expressing the proliferation marker Ki67 was between 1 in 2800 and 1 in 30,000.49 It was subsequently shown that the type B SLCs, the synovial fibroblasts, proliferated in situ,22,50 a finding consistent with the concept that type A synovial cells are macrophages. Mitotic activity of SLCs also is low in inflammatory conditions, such as rheuma­ toid arthritis, a condition associated with SLC hyperplasia. Coulton and coworkers51 reported “a few” mitotic figures in only 1 of 600 cases of rheumatoid arthritis synovium sam­ ples analyzed. Apart from the knowledge that synovial fibroblasts pro­ liferate slowly, little is known about their natural life span, recruitment, or mode of death. Apoptosis likely is involved in maintaining synovial homeostasis, but there is little in the literature on this subject. The dearth of information is likely to be explained by the lack of normal synovium avail­ able for analysis, in addition to the difficulty encountered when quantifying this process on histological sections owing to the rapid clearance of apoptotic bodies.52 Origin of Synovial Lining Cells There is little doubt that the type A SLC population iden­ tified by Barland and associates4 is bone marrow derived and represents cells of the mononuclear phagocyte system. The studies conducted by Edwards53,54 proved informative when they exploited the Beige (bg) mouse, which harbors a homozygous mutation that confers the presence of giant lysosomes in macrophages. It was shown that normal mice, bone marrow depleted through irradiation, were rescued



with bone marrow cells obtained from the bg mouse. Elec­ tron microscopic analysis of the synovium from the recipi­ ent animals revealed that type A SLCs contained the giant lysosomes of the donor bg mouse, and that these structures were never identified in type B cells. These findings pro­ vided powerful evidence that the type A SLCs represent macrophages, that they are recruited from the bone mar­ row, and that they were unrelated histogenetically to type B SLCs. In addition to immunohistochemistry, several other lines of evidence have added weight to the concept that type A SLCs are recruited from the bone marrow: (1) The op/op mouse, a spontaneously occurring mutant that fails to produce macrophage colony-stimulating factor because of a missense mutation in the csf-1 gene,55-57 has low numbers of circulating and resident macrophage colony-­stimulating factor–dependent macrophages, including those in the synovium. (2) Type A cells in rat synovium do not occur until after the development of synovial blood vessels.22 (3) Others have reported that type A SLCs were conspicu­ ous around vessels in the synovium in neonatal mice.6 (4) When synovial explants are placed in culture, the reduction in the type A SLCs is partially explained by their migration into the culture medium, an observation that reflects the process of migration of macrophages into the synovial fluid in vivo.1,58 (5) Macrophages are found around venules in disease states and constitute 80% of the intimal cells in inflammatory conditions, such as rheuma­ toid arthritis. Type B intimal cells represent a resident fibroblast popu­ lation in the synovial lining, but little is known about the cells from which they derive, and how their recruitment is regulated. The existence of a mesenchymal stem cell in the synovium is a prime candidate for the origin of the syno­ vial lining fibroblast, but this has not been substantiated. To date, a transcription factor directing mesenchymal stem cell differentiation into synovial fibroblast, similar to the factors required for commitment by this multipotential population into bone (cbfa-1), cartilage (Sox 9), and fat (peroxisome proliferator-activated receptor γ [PPARγ]), has not been identified.



PART 1 



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STRUCTURE AND FUNCTION OF BONE, JOINTS, AND CONNECTIVE TISSUE



SUBINTIMAL LAYER SLCs are not separated from the underlying subintima by a well-formed basement membrane composed of the typical trilaminar structure that is seen beneath epithelial mucosa elsewhere. Nevertheless, most components of basement membrane are present in the extracellular matrix surround­ ing SLCs. These components include tenascin X, perlecan (a heparin sulfate proteoglycan), collagen type 4, laminin, and fibrillin-1.59,60 Of note is the absence of laminin-5 and integrin α3β3γ2, which are components of epithelial hemidesmosomes.61 The subintima is composed of loose connective tissue of variable thickness and variable proportions of fibrous/col­ lagenous and adipose tissue depending on the anatomic site. Under normal healthy conditions, inflammatory cells are virtually absent from the subintima apart from a sprinkling of macrophages. A few mast cells also are present.62 Human synovial tissue also is a rich source of mesenchymal stem cells, and although it is unknown which compartment contains this cell population, some cells have the ability to self-renew, and differentiate into bone, cartilage, and fat in vitro, a phe­ nomenon that reflects its ability to regenerate in vivo.63-65



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There are three well-defined categories of subintima— the areolar, fibrous, and fatty/adipose types. Under the light microscope, areolar-type subintima, the most commonly studied, is generally found in larger joints where there is free movement (Fig. 2-5A). It is composed of fronds with a cel­ lular intimal lining and loose connective tissue in the sub­ intima, with little in the way of dense collagen fibers, and a rich vasculature. The fibrous subintima is composed of scant dense fibrous, poorly vascularized connective tissue and has an attenuated layer of SLCs (Fig. 2-5B). The adipose type contains abundant mature fat cells and has a single layer of SLCs. This is seen more commonly with aging and in intraarticular fat pads (Fig. 2-5C). The subintima contains collagen types I, III, V, and VI; glycosaminoglycans; proteoglycans; and extracellular matrices including tenascin and laminins. Integrin recep­ tors for collagens, laminin, and vitronectin are absent or at best weakly expressed by the subintimal cells. In con­ trast, receptors for fibronectin (CD49d and CD49e) are detected, and CD44, the HA receptor, is strongly expressed in most subintimal cells. β2 integrins are largely limited to perivascular areas, particularly in the subintimal zone, as is CD54.66



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Figure 2-5  Transmitted light photomicrographs of different morphologic types of synovial tissue. All photomicrographs show an intimal layer of one to two cells in depth. A, The areolar synovium is composed of villous fronds. Beneath the intimal lining layer, there is cellular loose fibrovascular fatty subintima. B, The fibrous synovium comprises dense collagenous material in the subintimal layer. C, The subintimal layer of the fatty synovial tissue is composed of less cellular mature adipose tissue with little collagen deposition.



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Subintimal Vasculature The vascular supply to the synovium is provided by many small vessels and is partly shared by the joint capsule, epiph­ yseal bone, and other perisynovial structures. Arteriovenous anastomoses communicate freely with the vascular supply to periosteum and to periarticular bone. As large synovial arteries enter the deep layers of the synovium near the cap­ sule, they branch, and branch again to form microvascular units in the more superficial subsynovial layers. Precapillary arterioles probably play a major role in controlling circula­ tion to the lining layer. The surface area of the synovial cap­ illary bed is large, and because it runs only a few cell layers deep to the surface, it has a role in trans-synovial exchange of molecules. Numerous physical factors influence synovial blood flow. Heat increases blood flow through synovial capillaries. Exer­ cise also increases synovial blood flow to normal joints, but may reduce the clearance rate of small molecules from the joint space. Experiments have shown a substantial vascu­ lar reserve capacity in normal articulations. Immobilization reduces synovial blood flow, and the pressure on synovial membrane from joint effusions can act to tamponade syno­ vial blood supply. The vascular endothelial lining cells express CD34 and CD31 (Fig. 2-6A). They also express receptors for the major components of basement membrane, including laminin and collagen IV, and the integrin receptors CD49a (laminin and collagen receptors), CD49d (fibronectin receptor), CD41, CD51 (vitronectin receptor), and CD61, the β3 integrin sub­ unit. Endothelial cells also express CD44, the HA receptor, and CD62, P-selectin, which acts as a receptor that supports binding of leukocytes to activated platelets and endothe­ lium. They are only weakly positive, however, for expres­ sion of CD54, intercellular adhesion molecule-1, an integral membrane protein of the immunoglobulin superfamily. The endothelial cells of capillaries in the superficial zone of the subintima are strongly positive for HLA-DR expression by immunohistochemistry, whereas cells in the larger vessels in the deep aspect of the membrane are negative.32,34 Subintimal Lymphatics Detailed analysis of the number and distribution of lym­ phatic vessels has been made possible with the use of the antibody to the lymphatic vessel endothelial HA ­receptor (LYVE-1) (Fig. 2-6B).67 This antibody is highly specific for lymphatic endothelial cells in lymphatic vessels and lymph node sinuses and does not react with endothelial cells of capillaries and other blood vessels that express CD34 and factor VIII–related antigen. The expression of LYVE-1 in lymphatic endothelial cells has been used as a marker to show that lymphatic vessels are less common in the fibrous synovium compared with the areolar and adipose variants of human subsynovial tissue. Detection of this molecule also reveals that lymphatics are present in the ­ superficial, intermediate, and deeper layers of synovial membrane from normal, osteoarthritic, and rheumatoid arthritic joints, although the number in the superficial subintimal layer is low in normal synovium. Little difference in the distribu­ tion and number is noted between normal and osteoarthritic synovium, where there is no villous ­hypertrophy. Lymphatic



channels are plentiful, however, in the subintimal layer in the presence of villous edema hypertrophy and chronic inflammation. Subintimal Nerve Supply The synovium has a rich network of sympathetic and sensory nerves. The former, which are myelinated and detected with the antibody against S100 protein, terminate close to blood vessels, where they regulate vascular tone (Fig. 2-6C, D, and E). The sensory nerves respond to proprioception and pain via large myelinated nerve fibers, and small (